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Contents and supplementary information for: Principles of
Gene Manipulation
Chapter 1
Chapter 2
Gene manipulation: an all-embracing technique
Basic techniques - (POGC02.pdf, 1,560KB)
Chapter 3
Cutting and joining DNA molecules
Chapter 4
Basic biology of plasmid and phage vectors
Chapter 5
Cosmids, phasmids and other advanced vectors
Chapter 6
Cloning strategies
Additional updated information on Cloning strategies
Chapter 7
Sequencing and mutagenesis
Chapter 8
Cloning in bacteria other than E. coli
Chapter 9
Cloning in Saccharomyces cerevisiae and other fungi
Chapter 10
Gene transfer to animal cells
Additional updated information on Gene transfer to animal
cells
Chapter 11
Genetic manipulation of animals
Additional updated information on Genetic manipulation of
animals
Chapter 12
Gene transfer to plants
Additional updated information on Gene transfer to plants
Chapter 13
Advances in transgenic technology
Additional updated information on Advances in transgenic
technology
Chapter 14
or (POGC13.pdf - size: 353KB)
Applications of recombinant DNA tecnology
Overview / Supplememtary Material / Related Titles / Related Websites / Ordering Information / Examination Copies / MCQ's /
New Edition
POGC01 9/11/2001 11:02 AM Page 1
CH A P T E R 1
Gene manipulation:
an all-embracing technique
Introduction
Sequence analysis
Occasionally technical developments in science
occur that enable leaps forward in our knowledge
and increase the potential for innovation. Molecular
biology and biomedical research experienced such
a revolutionary change in the mid-70s with the
development of gene manipulation. Although the
initial experiments generated much excitement, it is
unlikely that any of the early workers in the field
could have predicted the breadth of applications to
which the technique has been put. Nor could they
have envisaged that the methods they developed
would spawn an entire industry comprising several
hundred companies, of varying sizes, in the USA
alone.
The term gene manipulation can be applied to
a variety of sophisticated in vivo genetics as well as
to in vitro techniques. In fact, in most Western
countries there is a precise legal definition of gene
manipulation as a result of government legislation to control it. In the UK, gene manipulation is
defined as
the formation of new combinations of heritable
material by the insertion of nucleic acid molecules,
produced by whatever means outside the cell,
into any virus, bacterial plasmid or other vector
system so as to allow their incorporation into a
host organism in which they do not naturally
occur but in which they are capable of continued
propagation.
The definitions adopted by other countries are similar and all adequately describe the subject-matter
of this book. Simply put, gene manipulation permits stretches of DNA to be isolated from their host
organism and propagated in the same or a different
host, a technique known as cloning. The ability to
clone DNA has far-reaching consequences, as will
be shown below.
Cloning permits the isolation of discrete pieces of a
genome and their amplification. This in turn enables
the DNA to be sequenced. Analysis of the sequences
of some genetically well-characterized genes led to
the identification of the sequences and structures
which characterize the principal control elements
of gene expression, e.g. promoters, ribosome binding sites, etc. As this information built up it became
possible to scan new DNA sequences and identify
potential new genes, or open reading frames, because
they were bounded by characteristic motifs. Initially
this sequence analysis was done manually but to
the eye long runs of nucleotides have little meaning
and patterns evade recognition. Fortunately such
analyses have been facilitated by rapid increases
in the power of computers and improvements in software which have taken place contemporaneously
with advances in gene cloning. Now sequences can
be scanned quickly for a whole series of structural
features, e.g. restriction enzyme recognition sites,
start and stop signals for transcription, inverted
palindromes, sequence repeats, Z-DNA, etc., using
programs available on the Internet.
From the nucleotide sequence of a gene it is easy
to deduce the protein sequence which it encodes.
Unfortunately, we are unable to formulate a set of
general rules that allows us to predict a protein’s
three-dimensional structure from the amino acid
sequence of its polypeptide chain. However, based
on crystallographic data from over 300 proteins,
certain structural motifs can be predicted. Nor does
an amino acid sequence on its own give any clue to
function. The solution is to compare the amino acid
sequence with that of other better-characterized proteins: a high degree of homology suggests similarity
in function. Again, computers are of great value
since algorithms exist for comparing two sequences
or for comparing one sequence with a group of other
POGC01 9/11/2001 11:02 AM Page 2
2
CHAPTER 1
sequences simultaneously. The Internet has made
such comparisons easy because researchers can
access all the protein sequence data that are stored
in central databases, which are updated daily.
In vivo biochemistry
Any living cell, regardless of its origin, carries out a
plethora of biochemical reactions. To analyse these
different reactions, biochemists break open cells,
isolate the key components of interest and measure
their levels. They purify these components and try
to determine their performance characteristics. For
example, in the case of an enzyme, they might determine its substrate specificity and kinetic parameters,
such as Km and Vmax, and identify inhibitors and their
mode of action. From these data they try to build up
a picture of what happens inside the cell. However,
the properties of a purified enzyme in a test-tube
may bear little resemblance to its behaviour when
it shares the cell cytoplasm or a cell compartment
with thousands of other enzymes and chemical compounds. Understanding what happens inside cells
has been facilitated by the use of mutants. These
permit the determination of the consequences of
altered regulation or loss of a particular component or activity. Mutants have also been useful in
elucidating macromolecule structure and function.
However, the use of mutants is limited by the fact
that with classical technologies one usually has
little control over the type of mutant isolated and/or
location of the mutation.
Gene cloning provides elegant solutions to the above
problems. Once isolated, entire genes or groups of
genes can be introduced back into the cell type whence
they came or into different cell types or completely new
organisms, e.g. bacterial genes in plants or animals.
The levels of gene expression can be measured directly
or through the use of reporter molecules and can
be modulated up or down at the whim of the experimenter. Also, specific mutations, ranging from a
single base-pair to large deletions or additions, can
be built into the gene at any position to permit all
kinds of structural and functional analyses. Function
in different cell types can also be analysed, e.g. do
those structural features of a protein which result in
its secretion from a yeast cell enable it to be exported
from bacteria or higher eukaryotes? Experiments like
these permit comparative studies of macromolecular processes and, in some cases, gene cloning and
sequencing provides the only way to begin to understand such events as mitosis, cell division, telomere
structure, intron splicing, etc. Again, the Internet has
made such comparisons easy because researchers can
access all the protein sequence data that are stored
in central databases, which are updated daily.
The original goal of sequencing was to determine
the precise order of nucleotides in a gene. Then the
goal became the sequence of a small genome. First it
was that of a small virus (φX174, 5386 nucleotides).
Then the goal was larger plasmid and viral genomes,
then chromosomes and microbial genomes until
ultimately the complete genomes of higher eukaryotes
(humans, Arabidopsis) were sequenced (Table 1.1).
Table 1.1 Increases in sizes of genomes sequenced.
Genome sequenced
Year
Genome size
Bacteriophage fX174
Plasmid pBR322
Bacteriophage l
Epstein–Barr virus
Yeast chromosome III
Haemophilus influenzae
Saccharomyces cerevisiae
Ceanorhabditis elegans
Drosophila melanogaster
Homo sapiens
Arabidopsis thaliana
1977
1979
1982
1984
1992
1995
1996
1998
2000
2000
2000
5.38 kb
4.3 kb
48.5 kb
172 kb
315 kb
1.8 Mb
12 Mb
97 Mb
165 Mb
3000 Mb
125 Mb
Comment
First genome sequenced
First plasmid sequenced
First chromosome sequenced
First genome of cellular organism to be sequenced
First eukaryotic genome to be sequenced
First genome of multicellular organism to be sequenced
First mammalian genome to be sequenced
First plant genome to be sequenced
POGC01 9/11/2001 11:02 AM Page 3
Gene manipulation
Now the sequencing of large genomes has become
routine, albeit in specialist laboratories. Having the
complete genome sequence of an organism provides
us with fascinating insights into certain aspects
of its biology. For example, we can determine the
metabolic capabilities of a new microbe without
knowing anything about its physiology. However,
there are many aspects of cellular biology that cannot be ascertained from sequence data alone. For
example, what RNA species are made when in the
cell or organism life cycle and how fast do they
turn over? What proteins are made when and how
do the different proteins in a cell interact? How does
environment affect gene expression? The answers to
these questions are being provided by the new disciplines of genomics, proteomics and environomics
which rely heavily on the techniques of gene manipulation, which are discussed in later chapters. A
detailed presentation of whole-genome sequencing,
genomics and proteomics can be found in Primrose
and Twyman (2002).
3
recombinant DNA technology been greater than on
the practice of medicine.
The first medical benefit to arise from recombinant
DNA technology was the availability of significant
quantities of therapeutic proteins, such as human
growth hormone (HGH). This protein is used to treat
adolescents suffering from pituitary dwarfism to enable
them to achieve a normal height. Originally HGH was
purified from pituitary glands removed from cadavers.
However, a very large number of pituitary glands
are required to produce sufficient HGH to treat just
one child. Furthermore, some children treated with
pituitary-derived HGH have developed Creutzfeld–
Jakob syndrome. Following the cloning and expression of the HGH gene in Escherichia coli, it is possible
to produce enough HGH in a 10 litre fermenter to
treat hundreds of children. Since then, many different therapeutic proteins have become available for
the first time. Many of these proteins are also manufactured in E. coli but others are made in yeast or
animal cells and some in plants or the milk of animals.
The only common factor is that the relevant gene
has been cloned and overexpressed using the techniques of gene manipulation.
Medicine has benefited from recombinant DNA
technology in other ways (Fig. 1.1). New routes to
vaccines have been developed. The current hepatitis
B vaccine is based on the expression of a viral antigen on the surface of yeast cells and a recombinant
vaccine has been used to eliminate rabies from foxes
in a large part of Europe. Gene manipulation can
The new medicine
The developments in gene manipulation that have
taken place in the last 25 years have revolutionized
the study of biology. There is no subject area within
biology where recombinant DNA is not being used
and as a result the old divisions between subject areas
such as botany, genetics, zoology, biochemistry, etc.
are fast breaking down. Nowhere has the impact of
Profiling
Cloned P450s
Genetic
disease
Infectious
disease
Animal models
or human disease
Pharamacogenomics
Diagnostic
nucleic
acids
Plants
Therapeutic
small molecules
Therapeutic
nucleic
acids
MEDICINE
Microbes
Diagnostic
proteins
DNA
Vaccines
Therapeutic
proteins
Vaccines
Microbes
Microbes
Animals
Fig. 1.1 The impact of gene manipulation on the practice of medicine.
Plants
Gene therapy
Gene repair
Anti-sense drugs
POGC01 9/11/2001 11:02 AM Page 4
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CHAPTER 1
also be used to increase the levels of small molecules
within microbial cells. This can be done by cloning
all the genes for a particular biosynthetic pathway
and overexpressing them. Alternatively, it is possible to shut down particular metabolic pathways
and thus redirect particular intermediates towards
the desired end-product. This approach has been
used to facilitate production of chiral intermediates
and antibiotics. Novel antibiotics can also be created
by mixing and matching genes from organisms producing different but related molecules in a technique
known as combinatorial biosynthesis.
Gene cloning enables nucleic acid probes to be
produced readily and such probes have many uses
in medicine. For example, they can be used to
determine or confirm the identity of a microbial
pathogen or to diagnose pre- or perinatally an
inherited genetic disease. Increasingly, probes are
being used to determine the likelihood of adverse
reactions to drugs or to select the best class of drug
to treat a particular illness (pharmacogenomics). A
variant of this technique is to use cloned cytochrome
P450s to determine how a new drug will be metabolized and if any potentially toxic by-products will
result.
Nucleic acids are also being used as therapeutic
entities in their own right. For example, antisense
nucleic acids are being used to down-regulate gene
expression in certain diseases. In other cases, nucleic
acids are being administered to correct or repair
inherited gene defects (gene therapy/gene repair)
or as vaccines. In the reverse of gene repair, animals
are being generated that have mutations identical to those found in human disease. Note that
the use of antisense nucleic acids and gene therapy/
repair depends on the availability of information
on the exact cause of a disease. For most medical
conditions such information is lacking and currently available drugs are used to treat symptoms.
This situation will change significantly in the next
decade.
Biotechnology: the new industry
The early successes in overproducing mammalian
proteins in E. coli suggested to a few entrepreneurial
individuals that a new company should be formed to
exploit the potential of recombinant DNA technology.
Thus was Genentech born (Box 1.1). Since then
thousands of biotechnology companies have been
formed worldwide. As soon as major new developments in the science of gene manipulation are
reported, a rash of new companies are formed to
commercialize the new technology. For example,
many recently formed companies are hoping the
data from the Human Genome Sequencing Project
will result in the identification of a large number
of new proteins with potential for human therapy.
Others are using gene manipulation to understand
the regulation of transcription of particular genes,
arguing that it would make better therapeutic sense
to modulate the process with low-molecular-weight,
orally active drugs.
Although there are thousands of biotechnology companies, fewer than 100 have sales of their
products and even fewer are profitable. Already
many biotechnology companies have failed, but
the technology advances at such a rate that there
is no shortage of new company start-ups to take
their place. One group of biotechnology companies
that has prospered is those supplying specialist
reagents to laboratory workers engaged in gene
manipulation. In the very beginning, researchers
had to make their own restriction enzymes and
this restricted the technology to those with protein chemistry skills. Soon a number of companies were formed which catered to the needs of
researchers by supplying high-quality enzymes for
DNA manipulation. Despite the availability of these
enzymes, many people had great difficulty in cloning DNA. The reason for this was the need for careful quality control of all the components used in
the preparation of reagents, something researchers
are not good at! The supply companies responded
by making easy-to-use cloning kits in addition to
enzymes. Today, these supply companies can provide almost everything that is needed to clone,
express and analyse DNA and have thereby accelerated the use of recombinant DNA technology
in all biological disciplines. In the early days of
recombinant DNA technology, the development of
methodology was an end in itself for many academic
researchers. This is no longer true. The researchers
have gone back to using the tools to further our
POGC01 9/11/2001 11:02 AM Page 5
Gene manipulation
5
Box 1.1 The birth of an industry
Biotechnology is not new. Cheese, bread and yoghurt
are products of biotechnology and have been known
for centuries. However, the stock-market excitement
about biotechnology stems from the potential of
gene manipulation, which is the subject of this book.
The birth of this modern version of biotechnology
can be traced to the founding of the company
Genentech.
In 1976, a 27-year-old venture capitalist called
Robert Swanson had a discussion over a few beers
with a University of California professor, Herb Boyer.
The discussion centred on the commercial potential
of gene manipulation. Swanson’s enthusiasm for the
technology and his faith in it was contagious. By
the close of the meeting the decision was taken to
found Genentech (Genetic Engineering Technology).
Though Swanson and Boyer faced scepticism from
both the academic and business communities they
forged ahead with their idea. Successes came thick
and fast (see Table B1.1) and within a few years
they had proved their detractors wrong. Over
1000 biotechnology companies have been set up in
the USA alone since the founding of Genentech
but very, very few have been as successful.
Table B1.1 Key events at Genentech.
1976
1977
1978
1979
1980
1982
1984
1985
1987
1990
1990
Genentech founded
Genentech produced first human protein (somatostatin) in a microorganism
Human insulin cloned by Genentech scientists
Human growth hormone cloned by Genentech scientists
Genentech went public, raising $35 million
First recombinant DNA drug (human insulin) marketed (Genentech product licensed to Eli Lilly & Co.)
First laboratory production of factor VIII for therapy of haemophilia. Licence granted to Cutter Biological
Genentech launched its first product, Protropin (human growth hormone), for growth hormone deficiency in children
Genentech launched Activase (tissue plasminogen activator) for dissolving blood clots in heart-attack patients
Genentech launched Actimmune (interferon-g1b ) for treatment of chronic granulomatous disease
Genentech and the Swiss pharmaceutical company Roche complete a $2.1 billion merger
knowledge of biology, and the development of
new methodologies has largely fallen to the supply
companies.
The central role of E. coli
E. coli has always been a popular model system
for molecular geneticists. Prior to the development
of recombinant DNA technology, there existed a
large number of well-characterized mutants, gene
regulation was understood and there was a ready
availability of a wide selection of plasmids. Compared with other microbial systems it was matchless. It is not surprising, therefore, that the first
cloning experiments were undertaken in E. coli.
Subsequently, cloning techniques were extended to
a range of other microorganisms, such as Bacillus
subtilis, Pseudomonas sp., yeasts and filamentous
fungi, and then to higher eukaryotes. Curiously,
cloning in E. coli is technically easier than in any
other organism. As a result, it is rare for researchers
to clone DNA directly in other organisms. Rather,
DNA from the organism of choice is first manipulated in E. coli and subsequently transferred back
to the original host. Without the ability to clone
and manipulate DNA in E. coli, the application of
recombinant DNA technology to other organisms
would be greatly hindered.
POGC01 9/11/2001 11:02 AM Page 6
6
CHAPTER 1
Basic
Techniques
Cutting &
Joining DNA
Vectors
The role of vectors
Chapter 2
Agarose gel electrophoresis
Blotting (DNA, RNA, protein)
Nucleic acid hybridization
DNA transformation & electroporation
Polymerase chain reaction (PCR)
Restriction enzymes
Methods of joining DNA
Chapter 3
Basic properties of plasmids
Chapters 4 & 5
Desirable properties of vectors
Plasmids as vectors
Bacteriophage λ vectors
Single-stranded DNA vectors
Vectors for cloning large DNA molecules
Specialist vectors
Over-producing proteins
Cloning strategies
Cloning genomic DNA
cDNA cloning
Screening strategies
Expression cloning
Difference cloning
Chapter 6
Putting it
all together:
Cloning in
Practice
Analysing &
Changing
Cloned
Genes
Basic DNA sequencing
Analysing sequence data
Site-directed mutagenesis
Phage display
Chapter 7
Outline of the rest of the book
As noted above, E. coli has an essential role in recombinant DNA technology. Therefore, the first half of
the book is devoted to the methodology for manipulating genes in this organism (Fig. 1.2). Chapter 2
covers many of the techniques that are common
to all cloning experiments and are fundamental to
the success of the technology. Chapter 3 is devoted
to methods for selectively cutting DNA molecules
into fragments that can be readily joined together
again. Without the ability to do this, there would
be no recombinant DNA technology. If fragments
of DNA are inserted into cells, they fail to replicate
except in those rare cases where they integrate into
the chromosome. To enable such fragments to be
propagated, they are inserted into DNA molecules
(vectors) that are capable of extrachromosomal replication. These vectors are derived from plasmids
and bacteriophages and their basic properties are
described in Chapter 4. Originally, the purpose of
Fig. 1.2 ‘Roadmap’ outlining the basic
techniques in gene manipulation and
their relationships.
vectors was the propagation of cloned DNA but
today vectors fulfil many other roles, such as facilitating DNA sequencing, promoting expression of
cloned genes, facilitating purification of cloned gene
products, etc. The specialist vectors for these tasks
are described in Chapter 5. With this background in
place it is possible to describe in detail how to clone
the particular DNA sequences that one wants. There
are two basic strategies. Either one clones all the
DNA from an organism and then selects the very
small number of clones of interest or one amplifies
the DNA sequences of interest and then clones these.
Both these strategies are described in Chapter 6.
Once the DNA of interest has been cloned, it can
be sequenced and this will yield information on
the proteins that are encoded and any regulatory
signals that are present. There might also be a wish
to modify the DNA and/or protein sequence and
determine the biological effects of such changes.
The techniques for sequencing and changing cloned
genes are described in Chapter 7.
POGC01 9/11/2001 11:02 AM Page 7
Gene manipulation
Fig. 1.3 ‘Roadmap’ of the advanced
techniques in gene manipulation and
their application to organisms other
than E. coli.
7
Cloning in
Bacteria
Other Than
E.coli
Getting DNA into bacteria
Cloning in Gram-negative bacteria
Cloning in Gram-positive bacteria
Chapter 8
Why clone in fungi
Vectors for use in fungi
Expression of cloned DNA
Two hybrid system
Analysis of the whole genome
Chapter 9
Cloning in
Yeast &
Other
Fungi
Transformation of animal cells
Use of non-replicating DNA
Replication vectors
Viral transduction
Chapter 10
Gene
Transfer
To Animal
Cells
Transgenic mice
Other transgenic mammals
Transgenic birds, fish, Xenopus
Transgenic invertebrates
Chapter 11
Genetic
Manipulation
of Animals
Handling plant cells
Agrobacterium-mediated transformation
Direct DNA transfer
Plant viruses as vectors
Chapter 12
Genetic
Manipulation
of Plants
Inducible expression systems
Site-specific recombination
Gene inhibition
Insertional mutagenesis
Gene tagging
Entrapment constructs
Chapter 13
Advanced
Techniques
for Gene
Manipulation
in Plant and
Animals
In the second half of the book the specialist techniques for cloning in organisms other than E. coli
are described (Fig. 1.3). Each of these chapters can
be read in isolation from the other chapters in this
section, provided that there is a thorough understanding of the material from the first half of the
book. Chapter 8 details the methods for cloning in
other bacteria. Originally it was thought that some
of these bacteria, e.g. B. subtilis, would usurp the
position of E. coli. This has not happened and gene
manipulation techniques are used simply to better
understand the biology of these bacteria. Chapter 9
focuses on cloning in fungi, although the emphasis
is on the yeast Saccharomyces cerevisiae. Fungi are
eukaryotes and are useful model systems for investigating topics such as meiosis and mitosis, control of
cell division, etc. Animal cells can be cultured like
microorganisms and the techniques for cloning in
them are described in Chapter 10. Chapters 11 and
12 are devoted to the intricacies of cloning in animal
and plant representatives of higher eukaryotes and
Chapter 13 covers some cutting-edge techniques for
these same systems.
The concluding chapter is a survey of the different applications of recombinant DNA technology that are being exploited by the biotechnology
industry. Rather than going through application
after application, we have opted to show the interplay of different technologies by focusing on six
themes:
• Nucleic acid sequences as diagnostic tools.
• New drugs and new therapies for genetic diseases.
• Combating infectious disease.
• Protein engineering.
• Metabolic engineering.
• Plant breeding in the twenty-first century.
By treating the topic in this way we have been able to
show the interplay between some of the basic techniques and the sophisticated analysis now possible
with genome sequence information.
POGC02 9/11/2001 11:01 AM Page 8
CHAPTER 2
Basic techniques
The initial impetus for gene manipulation in vitro
came about in the early 1970s with the simultaneous development of techniques for:
• genetic transformation of Escherichia coli;
• cutting and joining DNA molecules;
• monitoring the cutting and joining reactions.
In order to explain the significance of these developments we must first consider the essential requirements of a successful gene-manipulation procedure.
and in the absence of replication will be diluted out of
their host cells. It should be noted that, even if a DNA
molecule contains an origin of replication, this may
not function in a foreign host cell.
There is an additional, subsequent problem. If the
early experiments were to proceed, a method was
required for assessing the fate of the donor DNA. In
particular, in circumstances where the foreign DNA
was maintained because it had become integrated in
the host DNA, a method was required for mapping the
foreign DNA and the surrounding host sequences.
The basic problems
The solutions: basic techniques
Before the advent of modern gene-manipulation
methods there had been many early attempts at
transforming pro- and eukaryotic cells with foreign
DNA. But, in general, little progress could be made.
The reasons for this are as follows. Let us assume
that the exogenous DNA is taken up by the recipient
cells. There are then two basic difficulties. First,
where detection of uptake is dependent on gene
expression, failure could be due to lack of accurate
transcription or translation. Secondly, and more
importantly, the exogenous DNA may not be maintained in the transformed cells. If the exogenous
DNA is integrated into the host genome, there is no
problem. The exact mechanism whereby this integration occurs is not clear and it is usually a rare
event. However this occurs, the result is that the
foreign DNA sequence becomes incorporated into
the host cell’s genetic material and will subsequently
be propagated as part of that genome. If, however,
the exogenous DNA fails to be integrated, it will
probably be lost during subsequent multiplication of
the host cells. The reason for this is simple. In order
to be replicated, DNA molecules must contain an
origin of replication, and in bacteria and viruses there
is usually only one per genome. Such molecules are
called replicons. Fragments of DNA are not replicons
If fragments of DNA are not replicated, the obvious
solution is to attach them to a suitable replicon.
Such replicons are known as vectors or cloning
vehicles. Small plasmids and bacteriophages are the
most suitable vectors for they are replicons in their
own right, their maintenance does not necessarily
require integration into the host genome and their
DNA can be readily isolated in an intact form. The
different plasmids and phages which are used as
vectors are described in detail in Chapters 4 and 5.
Suffice it to say at this point that initially plasmids
and phages suitable as vectors were only found in E.
coli. An important consequence follows from the use
of a vector to carry the foreign DNA: simple methods
become available for purifying the vector molecule,
complete with its foreign DNA insert, from transformed host cells. Thus not only does the vector
provide the replicon function, but it also permits the
easy bulk preparation of the foreign DNA sequence,
free from host-cell DNA.
Composite molecules in which foreign DNA has
been inserted into a vector molecule are sometimes
called DNA chimeras because of their analogy with
the Chimaera of mythology – a creature with the
head of a lion, body of a goat and tail of a serpent.
The construction of such composite or artificial
Introduction
POGC02 9/11/2001 11:01 AM Page 9
Basic techniques
recombinant molecules has also been termed genetic
engineering or gene manipulation because of the potential for creating novel genetic combinations by
biochemical means. The process has also been termed
molecular cloning or gene cloning because a line of
genetically identical organisms, all of which contain
the composite molecule, can be propagated and grown
in bulk, hence amplifying the composite molecule
and any gene product whose synthesis it directs.
Although conceptually very simple, cloning of
a fragment of foreign, or passenger, or target DNA
in a vector demands that the following can be
accomplished.
• The vector DNA must be purified and cut open.
• The passenger DNA must be inserted into the
vector molecule to create the artificial recombinant.
DNA joining reactions must therefore be performed.
Methods for cutting and joining DNA molecules are
now so sophisticated that they warrant a chapter of
their own (Chapter 3).
• The cutting and joining reactions must be readily monitored. This is achieved by the use of gel
electrophoresis.
• Finally, the artificial recombinant must be transformed into E. coli or another host cell. Further details
on the use of gel electrophoresis and transformation
of E. coli are given in the next section. As we have
noted, the necessary techniques became available at
about the same time and quickly led to many cloning
experiments, the first of which were reported in
1972 ( Jackson et al. 1972, Lobban & Kaiser 1973).
Agarose gel electrophoresis
The progress of the first experiments on cutting and
joining of DNA molecules was monitored by velocity
sedimentation in sucrose gradients. However, this
has been entirely superseded by gel electrophoresis.
Gel electrophoresis is not only used as an analytical
method, it is routinely used preparatively for the
purification of specific DNA fragments. The gel is
composed of polyacrylamide or agarose. Agarose is
convenient for separating DNA fragments ranging
in size from a few hundred base pairs to about 20 kb
(Fig. 2.1). Polyacrylamide is preferred for smaller
DNA fragments.
The mechanism responsible for the separation
of DNA molecules by molecular weight during gel
9
kb pairs
21.226
7.421
5.804
5.643
4.878
3.530
–
+
Fig. 2.1 Electrophoresis of DNA in agarose gels. The direction
of migration is indicated by the arrow. DNA bands have been
visualized by soaking the gel in a solution of ethidium bromide
(see Fig. 2.3), which complexes with DNA by intercalating
between stacked base-pairs, and photographing the orange
fluorescence which results upon ultraviolet irradiation.
electrophoresis is not well understood (Holmes
& Stellwagen 1990). The migration of the DNA
molecules through the pores of the matrix must play
an important role in molecular-weight separations
since the electrophoretic mobility of DNA in free
solution is independent of molecular weight. An
agarose gel is a complex network of polymeric
molecules whose average pore size depends on the
buffer composition and the type and concentration
of agarose used. DNA movement through the gel
was originally thought to resemble the motion of a
snake (reptation). However, real-time fluorescence
microscopy of stained molecules undergoing electrophoresis has revealed more subtle dynamics
(Schwartz & Koval 1989, Smith et al. 1989). DNA
molecules display elastic behaviour by stretching in
the direction of the applied field and then contracting into dense balls. The larger the pore size of the
POGC02 9/11/2001 11:01 AM Page 10
CHAPTER 2
gel, the greater the ball of DNA which can pass
through and hence the larger the molecules which
can be separated. Once the globular volume of the
DNA molecule exceeds the pore size, the DNA
molecule can only pass through by reptation. This
occurs with molecules about 20 kb in size and it is
difficult to separate molecules larger than this without recourse to pulsed electrical fields.
In pulsed-field gel electrophoresis (PFGE) (Schwartz
& Cantor 1984) molecules as large as 10 Mb can be
separated in agarose gels. This is achieved by causing the DNA to periodically alter its direction of
migration by regular changes in the orientation of
the electric field with respect to the gel. With each
change in the electric-field orientation, the DNA
must realign its axis prior to migrating in the new
direction. Electric-field parameters, such as the
direction, intensity and duration of the electric field,
are set independently for each of the different fields
and are chosen so that the net migration of the DNA
is down the gel. The difference between the direction
of migration induced by each of the electric fields is
the reorientation angle and corresponds to the angle
that the DNA must turn as it changes its direction of
migration each time the fields are switched.
A major disadvantage of PFGE, as originally
described, is that the samples do not run in straight
lines. This makes subsequent analysis difficult. This
problem has been overcome by the development of
improved methods for alternating the electrical field.
The most popular of these is contour-clamped homogeneous electrical-field electrophoresis (CHEF) (Chu
et al. 1986). In early CHEF-type systems (Fig. 2.2)
the reorientation angle was fixed at 120°. However,
in newer systems, the reorientation angle can be
varied and it has been found that for whole-yeast
chromosomes the migration rate is much faster with
an angle of 106° (Birren et al. 1988). Fragments of
DNA as large as 200–300 kb are routinely handled
in genomics work and these can be separated in a
matter of hours using CHEF systems with a reorientation angle of 90° or less (Birren & Lai 1994).
Aaij and Borst (1972) showed that the migration
rates of the DNA molecules were inversely proportional to the logarithms of the molecular weights.
Subsequently, Southern (1979a,b) showed that
plotting fragment length or molecular weight
against the reciprocal of mobility gives a straight
A–
B–
Migration of DNA
10
120°
B+
A+
Fig. 2.2 Schematic representation of CHEF (contour-clamped
homogeneous electrical field) pulsed-field gel electrophoresis.
NH2
N⊕
H2 N
Br –
C2H5
Fig. 2.3 Ethidium bromide.
line over a wider range than the semilogarithmic
plot. In any event, gel electrophoresis is frequently
performed with marker DNA fragments of known
size, which allow accurate size determination of an
unknown DNA molecule by interpolation. A particular advantage of gel electrophoresis is that the
DNA bands can be readily detected at high sensitivity. The bands of DNA in the gel are stained with
the intercalating dye ethidium bromide (Fig. 2.3),
and as little as 0.05 µg of DNA in one band can be
detected as visible fluorescence when the gel is
illuminated with ultraviolet light.
In addition to resolving DNA fragments of different lengths, gel electrophoresis can be used to separate different molecular configurations of a DNA
molecule. Examples of this are given in Chapter 4
(see p. 44). Gel electrophoresis can also be used for
investigating protein–nucleic acid interactions in
the so-called gel retardation or band shift assay. It is
based on the observation that binding of a protein
to DNA fragments usually leads to a reduction in
POGC02 9/11/2001 11:01 AM Page 11
Basic techniques
electrophoretic mobility. The assay typically involves
the addition of protein to linear double-stranded DNA
fragments, separation of complex and naked DNA
by gel electrophoresis and visualization. A review of
the physical basis of electrophoretic mobility shifts and
their application is provided by Lane et al. (1992).
11
• blotting of nucleic acids from gels;
• dot and slot blotting;
• colony and plaque blotting.
Colony and plaque blotting are described in detail on
pp. 104–105 and dot and slot blotting in Chapter 14.
Southern blotting
Nucleic acid blotting
Nucleic acid labelling and hybridization on membranes have formed the basis for a range of experimental techniques central to recent advances in our
understanding of the organization and expression
of the genetic material. These techniques may be
applied in the isolation and quantification of specific
nucleic acid sequences and in the study of their
organization, intracellular localization, expression
and regulation. A variety of specific applications
includes the diagnosis of infectious and inherited
disease. Each of these topics is covered in depth in
subsequent chapters.
An overview of the steps involved in nucleic acid
blotting and membrane hybridization procedures is
shown in Fig. 2.4. Blotting describes the immobilization of sample nucleic acids on to a solid support,
generally nylon or nitrocellulose membranes. The
blotted nucleic acids are then used as ‘targets’ in
subsequent hybridization experiments. The main
blotting procedures are:
The original method of blotting was developed by
Southern (1975, 1979b) for detecting fragments in
an agarose gel that are complementary to a given
RNA or DNA sequence. In this procedure, referred to
as Southern blotting, the agarose gel is mounted on
a filter-paper wick which dips into a reservoir containing transfer buffer (Fig. 2.5). The hybridization
membrane is sandwiched between the gel and a
stack of paper towels (or other absorbent material),
which serves to draw the transfer buffer through the
gel by capillary action. The DNA molecules are carried out of the gel by the buffer flow and immobilized
on the membrane. Initially, the membrane material
used was nitrocellulose. The main drawback with
this membrane is its fragile nature. Supported nylon
membranes have since been developed which have
greater binding capacity for nucleic acids in addition
to high tensile strength.
For efficient Southern blotting, gel pretreatment is
important. Large DNA fragments (> 10 kb) require a
longer transfer time than short fragments. To allow
Immobilization of nucleic acids
• Southern blot
• Northern blot
• Dot blot
• Colony/plaque lift
Pre-hybridization
Labelled DNA
or RNA probe
Hybridization
Stringency washes
Fig. 2.4 Overview of nucleic acid
blotting and hybridization (reproduced
courtesy of Amersham Pharmacia
Biotech).
Detection
Removal of probe
prior to reprobing
POGC02 9/11/2001 11:01 AM Page 12
12
CHAPTER 2
Weight < 0.75 kg
Glass plate
Paper tissues
3 sheets filter paper
Membrane
Gel
Plastic tray
Fig. 2.5 A typical capillary blotting apparatus.
uniform transfer of a wide range of DNA fragment
sizes, the electrophoresed DNA is exposed to a short
depurination treatment (0.25 mol/l HCl) followed by
alkali. This shortens the DNA fragments by alkaline
hydrolysis at depurinated sites. It also denatures the
fragments prior to transfer, ensuring that they are in
the single-stranded state and accessible for probing.
Finally, the gel is equilibrated in neutralizing solution
prior to blotting. An alternative method uses positively charged nylon membranes, which remove the
need for extended gel pretreatment. With them the
DNA is transferred in native (non-denatured) form
and then alkali-denatured in situ on the membrane.
After transfer, the nucleic acid needs to be fixed to
the membrane and a number of methods are available. Oven baking at 80°C is the recommended
method for nitrocellulose membranes and this can
also be used with nylon membranes. Due to the
flammable nature of nitrocellulose, it is important
that it is baked in a vacuum oven. An alternative
fixation method utilizes ultraviolet cross-linking. It
is based on the formation of cross-links between a
small fraction of the thymine residues in the DNA
and positively charged amino groups on the surface
of nylon membranes. A calibration experiment must
be performed to determine the optimal fixation period.
Following the fixation step, the membrane is placed
in a solution of labelled (radioactive or non-radioactive)
RNA, single-stranded DNA or oligodeoxynucleotide
which is complementary in sequence to the blottransferred DNA band or bands to be detected.
Conditions are chosen so that the labelled nucleic
acid hybridizes with the DNA on the membrane.
Since this labelled nucleic acid is used to detect and
locate the complementary sequence, it is called the
probe. Conditions are chosen which maximize the
rate of hybridization, compatible with a low background of non-specific binding on the membrane
(see Box 2.1). After the hybridization reaction has
been carried out, the membrane is washed to remove
unbound radioactivity and regions of hybridization
Box 2.1 Hybridization of nucleic acids on membranes
The hybridization of nucleic acids on membranes is a
widely used technique in gene manipulation and
analysis. Unlike solution hybridizations, membrane
hybridizations tend not to proceed to completion.
One reason for this is that some of the bound nucleic
acid is embedded in the membrane and is inaccessible
to the probe. Prolonged incubations may not generate
any significant increase in detection sensitivity.
The composition of the hybridization buffer can
greatly affect the speed of the reaction and the
sensitivity of detection. The key components of these
buffers are shown below:
Rate enhancers
Dextran sulphate and other polymers act as volume excluders to increase both the rate and the
extent of hybridization
Detergents and blocking agents
Dried milk, heparin and detergents such as sodium dodecyl sulphate (SDS) have been used
to depress non-specific binding of the probe to the membrane. Denhardt’s solution
(Denhardt 1966) uses Ficoll, polyvinylpyrrolidone and bovine serum albumin
Denaturants
Urea or formamide can be used to depress the melting temperature of the hybrid so that reduced
temperatures of hybridization can be used
Heterologous DNA
This can reduce non-specific binding of probes to non-homologous DNA on the blot
continued
POGC02 9/11/2001 11:01 AM Page 13
Basic techniques
13
Box 2.1 continued
Stringency control
Stringency can be regarded as the specificity with
which a particular target sequence is detected by
hybridization to a probe. Thus, at high stringency,
only completely complementary sequences will be
bound, whereas low-stringency conditions will allow
hybridization to partially matched sequences.
Stringency is most commonly controlled by the
temperature and salt concentration in the posthybridization washes, although these parameters
can also be utilized in the hybridization step.
In practice, the stringency washes are performed
under successively more stringent conditions
(lower salt or higher temperature) until the desired
result is obtained.
The melting temperature (Tm) of a probe–target
hybrid can be calculated to provide a starting-point
for the determination of correct stringency. The
Tm is the temperature at which the probe and
target are 50% dissociated. For probes longer than
100 base pairs:
Tm = 81.5°C + 16.6 log M + 0.41 (% G + C)
where M = ionic strength of buffer in moles/litre.
With long probes, the hybridization is usually carried
out at Tm − 25°C. When the probe is used to
detect partially matched sequences, the
hybridization temperature is reduced by 1°C
for every 1% sequence divergence between
probe and target.
Oligonucleotides can give a more rapid
hybridization rate than long probes as they can
be used at a higher molarity. Also, in situations
where target is in excess to the probe, for example
dot blots, the hybridization rate is diffusion-limited
and longer probes diffuse more slowly than
are detected autoradiographically by placing the
membrane in contact with X-ray film (see Box 2.2).
A common approach is to carry out the hybridization under conditions of relatively low stringency
which permit a high rate of hybridization, followed
by a series of post-hybridization washes of increasing
oligonucleotides. It is standard practice to use
oligonucleotides to analyse putative mutants
following a site-directed mutagenesis experiment
where the difference between parental and mutant
progeny is often only a single base-pair change
(see p. 132 et seq.).
The availability of the exact sequence of
oligonucleotides allows conditions for hybridization
and stringency washing to be tightly controlled so
that the probe will only remain hybridized when
it is 100% homologous to the target. Stringency is
commonly controlled by adjusting the temperature
of the wash buffer. The ‘Wallace rule’ (Lay Thein &
Wallace 1986) is used to determine the appropriate
stringency wash temperature:
Tm = 4 × (number of GC base pairs) + 2 × (number
of AT base pairs)
In filter hybridizations with oligonucleotide probes,
the hybridization step is usually performed at 5°C
below Tm for perfectly matched sequences. For every
mismatched base pair, a further 5°C reduction is
necessary to maintain hybrid stability.
The design of oligonucleotides for hybridization
experiments is critical to maximize hybridization
specificity. Consideration should be given to:
• probe length – the longer the oligonucleotide, the
less chance there is of it binding to sequences other
than the desired target sequence under conditions
of high stringency;
• oligonucleotide composition – the GC content
will influence the stability of the resultant hybrid
and hence the determination of the appropriate
stringency washing conditions. Also the presence
of any non-complementary bases will have an effect
on the hybridization conditions.
stringency (i.e. higher temperature or, more commonly, lower ionic strength). Autoradiography
following each washing stage will reveal any DNA
bands that are related to, but not perfectly complementary with, the probe and will also permit an
estimate of the degree of mismatching to be made.
POGC02 9/11/2001 11:01 AM Page 14
14
CHAPTER 2
Box 2.2 The principles of autoradiography
The localization and recording of a radiolabel within
a solid specimen is known as autoradiography
and involves the production of an image in a
photographic emulsion. Such emulsions consist of
silver halide crystals suspended in a clear phase
composed mainly of gelatin. When a b-particle or
g-ray from a radionuclide passes through the
emulsion, the silver ions are converted to silver atoms.
This results in a latent image being produced, which
is converted to a visible image when the image is
developed. Development is a system of amplification
in which the silver atoms cause the entire silver halide
crystal to be reduced to metallic silver. Unexposed
crystals are removed by dissolution in fixer, giving
an autoradiographic image which represents the
distribution of radiolabel in the original sample.
In direct autoradiography, the sample is placed
in intimate contact with the film and the radioactive
emissions produce black areas on the developed
autoradiograph. It is best suited to detection of
weak- to medium-strength b-emitting radionuclides
(3H, 14C, 35S). Direct autoradiography is not suited
to the detection of highly energetic b-particles,
such as those from 32P, or for g-rays emitted from
isotopes like 125I. These emissions pass through and
beyond the film, with the majority of the energy
being wasted. Both 32P and 125I are best detected
by indirect autoradiography.
Indirect autoradiography describes the technique
by which emitted energy is converted to light by
means of a scintillator, using fluorography or
intensifying screens. In fluorography the sample
is impregnated with a liquid scintillator. The
radioactive emissions transfer their energy to the
scintillator molecules, which then emit photons which
expose the photographic emulsion. Fluorography
is mostly used to improve the detection of weak
b-emitters (Fig. B2.1). Intensifying screens are
35S
+
3H
−
+
−
Fig. B2.1 Autoradiographs showing the detection of 35S- and 3H-labelled proteins in acrylamide gels with (+) and without
(−) fluorography. (Photo courtesy of Amersham Pharmacia Biotech.)
continued
POGC02 9/11/2001 11:01 AM Page 15
Basic techniques
15
Box 2.2 continued
sheets of a solid inorganic scintillator which are
placed behind the film. Any emissions passing
through the photographic emulsion are absorbed
by the screen and converted to light, effectively
superimposing a photographic image upon the
direct autoradiographic image.
The gain in sensitivity which is achieved by use of
indirect autoradiography is offset by non-linearity
of film response. A single hit by a b-particle or g-ray
can produce hundreds of silver atoms, but a single
hit by a photon of light produces only a single silver
atom. Although two or more silver atoms in a silver
halide crystal are stable, a single silver atom is
unstable and reverts to a silver ion very rapidly.
A
This means that the probability of a second photon
being captured before the first silver atom has
reverted is greater for large amounts of radioactivity
than for small amounts. Hence small amounts of
radioactivity are under-represented with the use of
fluorography and intensifying screens. This problem
can be overcome by a combination of pre-exposing a
film to an instantaneous flash of light (pre-flashing)
and exposing the autoradiograph at −70°C.
Pre-flashing provides many of the silver halide
crystals of the film with a stable pair of silver atoms.
Lowering the temperature to −70°C increases the
stability of a single silver atom, increasing the time
available to capture a second photon (Fig. B2.2).
B
C
Fig. B2.2 The improvement in sensitivity of detection of 125I-labelled IgG by autoradiography obtained by using an
intensifying screen and pre-flashed film. A, no screen and no pre-flashing; B, screen present but film not pre-flashed;
C, use of screen and pre-flashed film. (Photo courtesy of Amersham Pharmacia Biotech.)
POGC02 9/11/2001 11:01 AM Page 16
16
CHAPTER 2
Gene X
Restriction
endonuclease
Long DNA
fragments
Gel
electrophoresis
DNA
fragments
Short DNA
fragments
Genomic DNA
–
Genomic DNA
+
Agarose gel
(1) Denature in alkali
(2) Blot-transfer, bake
Autoradiography
(1) Hybridize nitrocellulose
with radioactive probe
Nitrocellulose
(2) Wash
Photographic
film
Images correspond only to
fragments containing gene X
sequences – estimate
fragment sizes from mobility
Radioactive RNA or
denatured DNA containing
sequences complementary
to gene X (radioactive probe)
The Southern blotting methodology can be extremely sensitive. It can be applied to mapping restriction sites around a single-copy gene sequence in a
complex genome such as that of humans (Fig. 2.6),
and when a ‘mini-satellite’ probe is used it can be
applied forensically to minute amounts of DNA (see
Chapter 14).
Northern blotting
Southern’s technique has been of enormous value,
but it was thought that it could not be applied
directly to the blot-transfer of RNAs separated by gel
electrophoresis, since RNA was found not to bind to
nitrocellulose. Alwine et al. (1979) therefore devised
a procedure in which RNA bands are blot-transferred
from the gel on to chemically reactive paper, where
they are bound covalently. The reactive paper is
prepared by diazotization of aminobenzyloxymethyl
paper (creating diazobenzyloxymethyl (DBM) paper),
which itself can be prepared from Whatman 540
paper by a series of uncomplicated reactions. Once
covalently bound, the RNA is available for hybrid-
Single stranded
DNA fragments
Fig. 2.6 Mapping restriction sites
around a hypothetical gene sequence
in total genomic DNA by the Southern
blot method.
Genomic DNA is cleaved with a
restriction endonuclease into hundreds
of thousands of fragments of various
sizes. The fragments are separated
according to size by gel electrophoresis
and blot-transferred on to nitrocellulose
paper. Highly radioactive RNA or
denatured DNA complementary in
sequence to gene X is applied to the
nitrocellulose paper bearing the blotted
DNA. The radiolabelled RNA or DNA
will hybridize with gene X sequences
and can be detected subsequently by
autoradiography, so enabling the sizes
of restriction fragments containing
gene X sequences to be estimated from
their electrophoretic mobility. By
using several restriction endonucleases
singly and in combination, a map of
restriction sites in and around gene
X can be built up.
ization with radiolabelled DNA probes. As before,
hybridizing bands are located by autoradiography.
Alwine et al.’s method thus extends that of Southern
and for this reason it has acquired the jargon term
northern blotting.
Subsequently it was found that RNA bands can
indeed be blotted on to nitrocellulose membranes
under appropriate conditions (Thomas 1980) and
suitable nylon membranes have been developed.
Because of the convenience of these more recent
methods, which do not require freshly activated paper,
the use of DBM paper has been superseded.
Western blotting
The term ‘western’ blotting (Burnette 1981) refers
to a procedure which does not directly involve nucleic
acids, but which is of importance in gene manipulation. It involves the transfer of electrophoresed
protein bands from a polyacrylamide gel on to a
membrane of nitrocellulose or nylon, to which they
bind strongly (Gershoni & Palade 1982, Renart &
Sandoval 1984). The bound proteins are then avail-
POGC02 9/11/2001 11:01 AM Page 17
Basic techniques
able for analysis by a variety of specific protein–ligand
interactions. Most commonly, antibodies are used to
detect specific antigens. Lectins have been used to
identify glycoproteins. In these cases the probe may
itself be labelled with radioactivity, or some other
‘tag’ may be employed. Often, however, the probe
is unlabelled and is itself detected in a ‘sandwich’
reaction, using a second molecule which is labelled,
for instance a species-specific second antibody, or
protein A of Staphylococcus aureus (which binds
to certain subclasses of IgG antibodies), or streptavidin (which binds to antibody probes that have
been biotinylated). These second molecules may
be labelled in a variety of ways with radioactive,
enzyme or fluorescent tags. An advantage of the
sandwich approach is that a single preparation of
labelled second molecule can be employed as a
general detector for different probes. For example,
an antiserum may be raised in rabbits which reacts
with a range of mouse immunoglobins. Such a
rabbit anti-mouse (RAM) antiserum may be radiolabelled and used in a number of different applications to identify polypeptide bands probed with
different, specific, monoclonal antibodies, each monoclonal antibody being of mouse origin. The sandwich method may also give a substantial increase
in sensitivity, owing to the multivalent binding of
antibody molecules.
Alternative blotting techniques
The original blotting technique employed capillary
blotting but nowadays the blotting is usually accomplished by electrophoretic transfer of polypeptides
from an SDS-polyacrylamide gel on to the membrane
(Towbin et al. 1979). Electrophoretic transfer is also
the method of choice for transferring DNA or RNA
from low-pore-size polyacrylamide gels. It can also
be used with agarose gels. However, in this case,
the rapid electrophoretic transfer process requires
high currents, which can lead to extensive heating
effects, resulting in distortion of agarose gels. The
use of an external cooling system is necessary to
prevent this.
Another alternative to capillary blotting is vacuumdriven blotting (Olszewska & Jones 1988), for which
several devices are commercially available. Vacuum
blotting has several advantages over capillary or
17
electrophoretic transfer methods: transfer is very
rapid and gel treatment can be performed in situ on
the vacuum apparatus. This ensures minimal gel
handling and, together with the rapid transfer, prevents significant DNA diffusion.
Transformation of E. coli
Early attempts to achieve transformation of E. coli
were unsuccessful and it was generally believed that
E. coli was refractory to transformation. However,
Mandel and Higa (1970) found that treatment with
CaC12 allowed E. coli cells to take up DNA from bacteriophage λ. A few years later Cohen et al. (1972)
showed that CaC12-treated E. coli cells are also effective recipients for plasmid DNA. Almost any strain of
E. coli can be transformed with plasmid DNA, albeit
with varying efficiency, whereas it was thought that
only recBC− mutants could be transformed with linear bacterial DNA (Cosloy & Oishi 1973). Later,
Hoekstra et al. (1980) showed that recBC+ cells can
be transformed with linear DNA, but the efficiency is
only 10% of that in otherwise isogenic recBC− cells.
Transformation of recBC− cells with linear DNA is
only possible if the cells are rendered recombinationproficient by the addition of a sbcA or sbcB mutation. The fact that the recBC gene product is an
exonuclease explains the difference in transformation efficiency of circular and linear DNA in recBC+
cells.
As will be seen from the next chapter, many bacteria contain restriction systems which can influence
the efficiency of transformation. Although the complete function of these restriction systems is not yet
known, one role they do play is the recognition and
degradation of foreign DNA. For this reason it is
usual to use a restriction-deficient strain of E. coli as
a transformable host.
Since transformation of E. coli is an essential step
in many cloning experiments, it is desirable that it be
as efficient as possible. Several groups of workers
have examined the factors affecting the efficiency of
transformation. It has been found that E. coli cells
and plasmid DNA interact productively in an environment of calcium ions and low temperature
(0–5°C), and that a subsequent heat shock (37–45°C)
is important, but not strictly required. Several other
factors, especially the inclusion of metal ions in
POGC02 9/11/2001 11:01 AM Page 18
18
CHAPTER 2
addition to calcium, have been shown to stimulate
the process.
A very simple, moderately efficient transformation procedure for use with E. coli involves resuspending log-phase cells in ice-cold 50 mmol/l
calcium chloride at about 1010 cells/ml and keeping
them on ice for about 30 min. Plasmid DNA (0. 1 µg)
is then added to a small aliquot (0.2 ml) of these now
competent (i.e. competent for transformation) cells,
and the incubation on ice continued for a further
30 min, followed by a heat shock of 2 min at 42°C.
The cells are then usually transferred to nutrient
medium and incubated for some time (30 min to
1 h) to allow phenotypic properties conferred by the
plasmid to be expressed, e.g. antibiotic resistance
commonly used as a selectable marker for plasmidcontaining cells. (This so-called phenotypic lag
may not need to be taken into consideration with
high-level ampicillin resistance. With this marker,
significant resistance builds up very rapidly, and
ampicillin exerts its effect on cell-wall biosynthesis
only in cells which have progressed into active
growth.) Finally the cells are plated out on selective
medium. Just why such a transformation procedure
is effective is not fully understood (Huang & Reusch
1995). The calcium chloride affects the cell wall and
may also be responsible for binding DNA to the cell
surface. The actual uptake of DNA is stimulated by
the brief heat shock.
Hanahan (1983) has re-examined factors that
affect the efficiency of transformation, and has devised
a set of conditions for optimal efficiency (expressed
as transformants per µg plasmid DNA) applicable to
most E. coli K12 strains. Typically, efficiencies of 107
to 109 transformants/µg can be achieved depending
on the strain of E. coli and the method used (Liu &
Rashidbaigi 1990). Ideally, one wishes to make a
large batch of competent cells and store them frozen
for future use. Unfortunately, competent cells made
by the Hanahan procedure rapidly lose their competence on storage. Inoue et al. (1990) have optimized
the conditions for the preparation of competent cells.
Not only could they store cells for up to 40 days at
−70°C while retaining efficiencies of 1–5 × 109 cfu/µg,
but competence was affected only minimally by salts
in the DNA preparation.
There are many enzymic activities in E. coli which
can destroy incoming DNA from non-homologous
sources (see Chapter 3) and reduce the transformation efficiency. Large DNAs transform less efficiently, on a molar basis, than small DNAs. Even with
such improved transformation procedures, certain
potential gene-cloning experiments requiring large
numbers of clones are not reliable. One approach
which can be used to circumvent the problem of low
transformation efficiencies is to package recombinant DNA into virus particles in vitro. A particular
form of this approach, the use of cosmids, is described
in detail in Chapter 5. Another approach is electroporation, which is described below.
Electroporation
A rapid and simple technique for introducing cloned
genes into a wide variety of microbial, plant and animal cells, including E. coli, is electroporation. This
technique depends on the original observation by
Zimmerman & Vienken (1983) that high-voltage
electric pulses can induce cell plasma membranes to
fuse. Subsequently it was found that, when subjected to electric shock, the cells take up exogenous
DNA from the suspending solution. A proportion of
these cells become stably transformed and can be
selected if a suitable marker gene is carried on the
transforming DNA. Many different factors affect the
efficiency of electroporation, including temperature,
various electric-field parameters (voltage, resistance
and capacitance), topological form of the DNA,
and various host-cell factors (genetic background,
growth conditions and post-pulse treatment). Some
of these factors have been reviewed by Hanahan
et al. (1991).
With E. coli, electroporation has been found to give
plasmid transformation efficiencies (109 cfu/µg DNA)
comparable with the best CaC12 methods (Dower et al.
1988). More recently, Zhu and Dean (1999) have
reported 10-fold higher transformation efficiencies
with plasmids (9 × 109 transformants/µg) by coprecipitating the DNA with transfer RNA (tRNA)
prior to electroporation. With conventional CaCl2mediated transformation, the efficiency falls off
rapidly as the size of the DNA molecule increases
and is almost negligible when the size exceeds 50 kb.
While size also affects the efficiency of electroporation
(Sheng et al. 1995), it is possible to get transformation efficiencies of 106 cfu/µg DNA with molecules
POGC02 9/11/2001 11:01 AM Page 19
Basic techniques
as big as 240 kb. Molecules three to four times this
size also can be electroporated successfully. This is
important because much of the work on mapping
and sequencing of genomes demands the ability
to handle large fragments of DNA (see p. 64 and
p. 126).
Transformation of other organisms
Although E. coli often remains the host organism of
choice for cloning experiments, many other hosts
are now used, and with them transformation may
still be a critical step. In the case of Gram-positive
bacteria, the two most important groups of organisms are Bacillus spp. and actinomycetes. That B.
subtilis is naturally competent for transformation
has been known for a long time and hence the genetics of this organism are fairly advanced. For this
reason B. subtilis is a particularly attractive alternative prokaryotic cloning host. The significant features
of transformation with this organism are detailed
in Chapter 8. Of particular relevance here is that
it is possible to transform protoplasts of B. subtilis, a
technique which leads to improved transformation
frequencies. A similar technique is used to transform
actinomycetes, and recently it has been shown that
the frequency can be increased considerably by first
entrapping the DNA in liposomes, which then fuse
with the host-cell membrane.
In later chapters we discuss ways, including electroporation, in which cloned DNA can be introduced
into eukaryotic cells. With animal cells there is no
great problem as only the membrane has to be
crossed. In the case of yeast, protoplasts are required
(Hinnen et al. 1978). With higher plants one strategy that has been adopted is either to package the
DNA in a plant virus or to use a bacterial plant
pathogen as the donor. It has also been shown that
protoplasts prepared from plant cells are competent
for transformation. A further remarkable approach
that has been demonstrated with plants and animals
(Klein & Fitzpatrick-McElligott 1993) is the use of
microprojectiles shot from a gun (p. 238).
Animal cells, and protoplasts of yeast, plant and
bacterial cells are susceptible to transformation by
liposomes (Deshayes et al. 1985). A simple transformation system has been developed which makes use of
liposomes prepared from a cationic lipid (Felgner
19
et al. 1987). Small unilamellar (single-bilayer) vesicles are produced. DNA in solution spontaneously
and efficiently complexes with these liposomes (in
contrast to previously employed liposome encapsidation procedures involving non-ionic lipids). The
positively charged liposomes not only complex with
DNA, but also bind to cultured animal cells and are
efficient in transforming them, probably by fusion
with the plasma membrane. The use of liposomes as
a transformation or transfection system is called
lipofection.
The polymerase chain reaction (PCR)
The impact of the PCR upon molecular biology has
been profound. The reaction is easily performed, and
leads to the amplification of specific DNA sequences
by an enormous factor. From a simple basic principle, many variations have been developed with
applications throughout gene technology (Erlich
1989, Innis et al. 1990). Very importantly, the PCR
has revolutionized prenatal diagnosis by allowing
tests to be performed using small samples of fetal tissue. In forensic science, the enormous sensitivity of
PCR-based procedures is exploited in DNA profiling;
following the publicity surrounding Jurassic Park,
virtually everyone is aware of potential applications in palaeontology and archaeology. Many other
processes have been described which should produce equivalent results to a PCR (for review, see
Landegran 1996) but as yet none has found widespread use.
In many applications of the PCR to gene manipulation, the enormous amplification is secondary
to the aim of altering the amplified sequence. This
often involves incorporating extra sequences at the
ends of the amplified DNA. In this section we shall
consider only the amplification process. The applications of the PCR will be described in appropriate places.
Basic reaction
First we need to consider the basic PCR. The
principle is illustrated in Fig. 2.7. The PCR involves
two oligonucleotide primers, 17–30 nucleotides in
length, which flank the DNA sequence that is to be
amplified. The primers hybridize to opposite strands
of the DNA after it has been denatured, and are
POGC02 9/11/2001 11:01 AM Page 20
20
CHAPTER 2
Cycle 1
5’+
3’–
Denaturation by
heat followed by
primer annealing
5’+
3’ Double stranded
5’ DNA target
3’
and
5’
3’–
3’
5’
3’
5’
DNA synthesis
(primer extension)
5’
3’
3’
and
5’
5’
3’
3’
5’
Denaturation by heat followed by primer
annealing and DNA synthesis
Cycle 2
5’
3’
3’
+
5’
3’
+
5’
3’
+
5’
3’
5’
5’
5’
3’
3’
3’
5’
Denaturation by heat followed by primer
annealing and DNA synthesis
Cycle 3
5’
3’
3’
3’
5’
5’
3’
3’
5’
5’
3’
3’
5’
5’
3’
5’
5’
3’
5’
5’
3’
3’
5’
5’
3’
5’
3’
3’
5’
3’
Repeated cycles lead to exponential
doubling of the target sequence
orientated so that DNA synthesis by the polymerase
proceeds through the region between the two
primers. The extension reactions create two doublestranded target regions, each of which can again be
denatured ready for a second cycle of hybridization
and extension. The third cycle produces two doublestranded molecules that comprise precisely the
target region in double-stranded form. By repeated
cycles of heat denaturation, primer hybridization
and extension, there follows a rapid exponential
accumulation of the specific target fragment of
DNA. After 22 cycles, an amplification of about 106fold is expected (Fig. 2.8), and amplifications of this
order are actually attained in practice.
In the original description of the PCR method
(Mullis & Faloona 1987, Saiki et al. 1988, Mullis
1990), Klenow DNA polymerase was used and,
because of the heat-denaturation step, fresh enzyme
had to be added during each cycle. A breakthrough
came with the introduction of Taq DNA polymerase
(Lawyer et al. 1989) from the thermophilic bacterium
Thermus aquaticus. The Taq DNA polymerase is
resistant to high temperatures and so does not need
to be replenished during the PCR (Erlich et al. 1988,
Sakai et al. 1988). Furthermore, by enabling the
extension reaction to be performed at higher temperatures, the specificity of the primer annealing is
not compromised. As a consequence of employing
the heat-resistant enzyme, the PCR could be automated very simply by placing the assembled reaction
in a heating block with a suitable thermal cycling
programme (see Box 2.3).
3’
5’
Fig. 2.7 (left) The polymerase chain reaction. In cycle 1 two
primers anneal to denatured DNA at opposite sides of the
target region, and are extended by DNA polymerase to give
new strands of variable length. In cycle 2, the original strands
and the new strands from cycle 1 are separated, yielding a
total of four primer sites with which primers anneal. The
primers that are hybridized to the new strands from cycle 1
are extended by polymerase as far as the end of the template,
leading to a precise copy of the target region. In cycle 3,
double-stranded DNA molecules are produced (highlighted in
colour) that are precisely identical to the target region.
Further cycles lead to exponential doubling of the target
region. The original DNA strands and the variably extended
strands become negligible after the exponential increase of
target fragments.
POGC02 9/11/2001 11:01 AM Page 21
Basic techniques
Cycle number
Number of double-stranded
target molecules
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
0
0
2
4
8
16
32
64
128
256
512
1024
2048
4096
8192
16,384
32,768
65,536
131,072
262,144
524,288
1,048,576
2,097,152
4,194,304
8,388,608
16,777,216
33,554,432
67,108,864
134,217,728
268,435,456
Fig. 2.8 Theoretical PCR amplification of a target fragment
with increasing number of cycles.
Recent developments have sought to minimize
amplification times. Such systems have used small
reaction volumes in glass capillaries to give large
surface area-to-volume ratios. This results in almost
instantaneous temperature equilibration and minimal
annealing and denaturation times. This, accompanied by temperature ramp rates of 10–20°C/s, made
possible by the use of turbulent forced hot-air systems to heat the sample, results in an amplification
reaction completed in tens of minutes.
While the PCR is simple in concept, practically
there are a large number of variables which can
influence the outcome of the reaction. This is especially important when the method is being used with
rare samples of starting material or if the end result
has diagnostic or forensic implications. For a detailed
analysis of the factors affecting the PCR, the reader
should consult McDowell (1999). There are many
substances present in natural samples (e.g. blood,
faeces, environmental materials) which can inter-
21
fere with the PCR, and ways of eliminating them
have been reviewed by Bickley and Hopkins (1999).
RT-PCR
The thermostable polymerase used in the basic PCR
requires a DNA template and hence is limited to the
amplification of DNA samples. There are numerous
instances in which the amplification of RNA would
be preferred. For example, in analyses involving the
diffierential expression of genes in tissues during
development or the cloning of DNA derived from an
mRNA (complementary DNA or cDNA), particularly
a rare mRNA. In order to apply PCR methodology
to the study of RNA, the RNA sample must first be
reverse-transcribed to cDNA to provide the necessary
DNA template for the thermostable polymerase. This
process is called reverse transcription (RT), hence
the name RT-PCR.
Avian myeloblastosis virus (AMV) or Moloney
murine leukaemia virus (MuLV) reverse transcriptases are generally used to produce a DNA copy of
the RNA template. Various strategies can be adopted
for first-strand cDNA synthesis (Fig. 2.9).
Long accurate PCR (LA-PCR)
Amplification of long DNA fragments is desirable for
numerous applications of gene manipulation. The
basic PCR works well when small fragments are
amplified. The efficiency of amplification and therefore the yield of amplified fragments decrease significantly as the size of the amplicon increases over 5 kb.
This decrease in yield of longer amplified fragments
is attributable to partial synthesis across the desired
sequence, which is not a suitable substrate for the
subsequent cycles. This is demonstrated by the presence of smeared, as opposed to discrete, bands on a gel.
Barnes (1994) and Cheng et al. (1994) examined
the factors affecting the thermostable polymerization across larger regions of DNA and identified key
variables affecting the yield of longer PCR fragments. Most significant of these was the absence of
a 3′–5′ exonuclease (proofreading) activity in Taq
polymerase. Presumably, when the Taq polymerase
misincorporates a dNTP, subsequent extension of
the strand either proceeds very slowly or stops
completely. To overcome this problem, a second
POGC02 9/11/2001 11:01 AM Page 22
22
CHAPTER 2
Box 2.3 The polymerase chain reaction achieves enormous amplifications,
of specific target sequence, very simply
The reaction is assembled in a single tube, and then
placed in a thermal cycler (a programmable
heating/cooling block), as described below.
A typical PCR for amplifying a human genomic
DNA sequence has the following composition. The
reaction volume is 100 ml.
Input genomic DNA, 0.1–1 mg
Primer 1, 20 pmol
Primer 2, 20 pmol
20 mmol/l Tris-HCl, pH 8.3 (at 20°C)
1.5 mmol/l magnesium chloride
25 mmol/l potassium chloride
50 mmol/l each deoxynucleoside triphosphate
(dATP, dCTP, dGTP, dTTP)
2 units Taq DNA polymerase
A layer of mineral oil is placed over the reaction
mix to prevent evaporation.
The reaction is cycled 25–35 times, with the
following temperature programme:
Denaturation
94°C, 0.5 min
Primer annealing 55°C,1.5 min
Extension
72°C, 1 min
Typically, the reaction takes some 2–3 h overall.
Notes:
• The optimal temperature for the annealing step
will depend upon the primers used.
• The pH of the Tris-HCl buffer decreases markedly
with increasing temperature. The actual pH varies
between about 6.8 and 7.8 during the thermal cycle.
• The time taken for each cycle is considerably
longer than 3 min (0.5 + 1.5 + 1 min), depending
upon the rates of heating and cooling between steps,
but can be reduced considerably by using turbo
systems (p. 21).
• The standard PCR does not efficiently amplify
sequences much longer than about 3 kb.
Random primer
5’
A A A A A A A 3’
mRNA
1st strand cDNA
A A A A A A A 3’
T T T T T T T 5’
mRNA
1st strand cDNA
A A A A A A A 3’
mRNA
1st strand cDNA
3’
random primer
Oligo (dT) primer
5’
3’
Sequence-specific primer
5’
3’
primer
thermostable polymerase with proofreading capability is added. Thermostable DNA polymerases with
proofreading capabilities are listed in Table 2.1.
Key factors affecting the PCR
The specificity of the PCR depends crucially upon the
primers. The following factors are important in
choosing effective primers.
Fig. 2.9 Three strategies for synthesis of
first-strand cDNA. (a) Random primer;
(b) oligo (dT) primer; (c) sequence-specific
primer.
• Primers should be 17 to 30 nucleotides in length.
• A GC content of about 50% is ideal. For primers with a low GC content, it is desirable to
choose a long primer so as to avoid a low melting
temperature.
• Sequences with long runs (i.e. more than three or
four) of a single nucleotide should be avoided.
• Primers with significant secondary structure are
undesirable.
POGC02 9/11/2001 11:01 AM Page 23
Basic techniques
Table 2.1 Sources of thermostable DNA polymerases
with proofreading (3′–5′ exonuclease) activity.
DNA polymerase
Tma
Deep VentTM
Tli
Pfu
Pwo
Source
Thermotoga maritima
Pyrococcus sp.
Thermococcus litoralis
Pyrococcus furiosus
Pyrococcus woesi
• There should be no complementarity between the
two primers. The great majority of primers which
conform with these guidelines can be made to work,
although not all comparable primer sets are equally
effective even under optimized conditions.
In carrying out a PCR it is usual to employ a
hot-start protocol. This entails adding the DNA
polymerase after the heat-denaturation step of the
first cycle, the addition taking place at a temperature
at or above the annealing temperature and just prior
to the annealing step of the first cycle. The hot start
overcomes the problem that would arise if the DNA
polymerase were added to complete the assembly
of the PCR reaction mixture at a relatively low
temperature. At low temperature, below the desired
hybridization temperature for the primer (typically
in the region 45–60°C), mismatched primers will
form and may be extended somewhat by the polymerase. Once extended, the mismatched primer is
stabilized at the unintended position. Having been
incorporated into the extended DNA during the
first cycle, the primer will hybridize efficiently in
subsequent cycles and hence may cause the amplification of a spurious product.
Alternatives to the hot-start protocol include the
use of Taq polymerase antibodies, which are inactivated as the temperature rises (Taylor & Logan 1995),
and AmpliTaq GoldTM, a modified Taq polymerase
that is inactive until heated to 95°C (Birch 1996).
Yet another means of inactivating Taq DNA
polymerase at ambient temperatures is the SELEX
method (systematic evolution of ligands by exponential enrichment). Here the polymerase is
reversibly inactivated by the binding of nanomolar
amounts of a 70-mer, which is itself a poor poly-
23
merase substrate and should not interfere with the
amplification primers (Dang & Jayasena 1996).
In order to minimize further the amplification of
spurious products, the strategy of nested primers may
be deployed. Here the products of an initial PCR
amplification are used to seed a second PCR amplification, in which one or both primers are located
internally with respect to the primers of the first
PCR. Since there is little chance of the spurious products containing sequences capable of hybridizing
with the second primer set, the PCR with these
nested primers selectively amplifies the sought-after
DNA.
As noted above, the Taq DNA polymerase lacks a
3′–5′ proofreading exonuclease. This lack appears
to contribute to errors during PCR amplification due
to misincorporation of nucleotides (Eckert & Kunkel
1990). Partly to overcome this problem, other
thermostable DNA polymerases with improved
fidelity have been sought, although the Taq DNA
polymerase remains the most widely used enzyme
for PCR. In certain applications, especially where
amplified DNA is cloned, it is important to check the
nucleotide sequence of the cloned product to reveal
any mutations that may have occurred during the
PCR. The fidelity of the amplification reaction can be
assessed by cloning, sequencing and comparing
several independently amplified molecules.
Real-time quantitative PCR
There are many applications of the PCR where it
would be advantageous to be able to quantify the
amount of starting material. Theoretically, there is
a quantitative relationship between the amount of
starting material (target sequence) and the amount
of PCR product at any given cycle. In practice,
replicate reactions yield different amounts of product, making quantitation unreliable. Higuchi et al.
(1992, 1993) pioneered the use of ethidium bromide
to quantify PCR products as they accumulate. Amplification produces increasing amounts of doublestranded DNA, which binds ethidium bromide,
resulting in an increase in fluorescence. By plotting
the increase in fluorescence versus cycle number it is
possible to analyse the PCR kinetics in real time. This
is much more satisfactory than analysing product
accumulation after a fixed number of cycles.
POGC02 9/11/2001 11:01 AM Page 24
24
CHAPTER 2
Reporter
5’
3’
5’
Forward
primer
R
Quencher
Probe
Q
Reverse
primer
R
Binding of
primers and probe
5’
3’
5’
Polymerization
5’
3’
5’
Strand
displacement
5’
3’
5’
Release of
reporter
5’
3’
5’
Polymerization
complete
Q
5’
3’
5’
R
Q
5’
3’
5’
R
Q
5’
3’
5’
R
5’
3’
5’
5’
3’
5’
Q
The principal drawback to the use of ethidium
bromide is that both specific and non-specific products generate a signal. This can be overcome by the
use of probe-based methods for assaying product
accumulation (Livak et al. 1995). The probes used
are oligonucleotides with a reporter fluorescent dye
attached at the 5′ end and a quencher dye at the 3′
end. While the probe is intact, the proximity of
the quencher reduces the fluorescence emitted by
the reporter dye. If the target sequence is present, the
probe anneals downstream from one of the primer
sites. As the primer is extended, the probe is cleaved
by the 5′ nuclease activity of the Taq polymerase
Fig. 2.10 Real-time quantitative PCR. See
text for details.
(Fig. 2.10). This cleavage of the probe separates
the reporter dye from the quencher dye, thereby
increasing the reporter-dye signal. Cleavage removes
the probe from the target strand, allowing primer
extension to continue to the end of the template
strand. Additional reporter-dye molecules are cleaved
from their respective probes with each cycle, effecting an increase in fluorescence intensity proportional
to the amount of amplicon produced.
Instrumentation has been developed which
combines thermal cycling with measurement of
fluorescence, thereby enabling the progress of the
PCR to be monitored in real time. This revolutionizes
POGC02 9/11/2001 11:01 AM Page 25
Basic techniques
the way one approaches PCR-based quantitation of
DNA. Reactions are characterized by the point in
time during cycling when amplification of a product
is first detected, rather than by the amount of PCR
product accumulated after a fixed number of cycles.
The higher the starting copy number of the target,
25
the sooner a significant increase in fluorescence is
noted. Quantitation of the amount of target in
unknown samples is achieved by preparing a standard curve, using different starting copy numbers of
the target sequence.
POGC03 9/11/2001 11:00 AM Page 26
CHAPTER 3
Cutting and joining DNA
molecules
Cutting DNA molecules
Before 1970 there was no method of cleaving DNA
at discrete points. All the available methods for
fragmenting DNA were non-specific. The available
endonucleases had little site specificity and chemical
methods produced very small fragments of DNA.
The only method where any degree of control could
be exercised was the use of mechanical shearing.
The long, thin threads which constitute duplex
DNA molecules are sufficiently rigid to be very easily
broken by shear forces in solution. Intense sonication with ultrasound can reduce the length to about
300 nucleotide pairs. More controlled shearing
can be achieved by high-speed stirring in a blender.
Typically, high-molecular-weight DNA is sheared
to a population of molecules with a mean size of
about 8 kb by stirring at 1500 rev/min for 30 min
(Wensink et al. 1974). Breakage occurs essentially
at random with respect to DNA sequence. The termini consist of short, single-stranded regions which
may have to be taken into account in subsequent
joining procedures.
During the 1960s, phage biologists elucidated the
biochemical basis of the phenomenon of host restriction and modification. The culmination of this work
was the purification of the restriction endonuclease
of Escherichia coli K12 by Meselson and Yuan (1968).
Since this endonuclease cuts unmodified DNA into
large discrete fragments, it was reasoned that it must
recognize a target sequence. This in turn raised the
prospect of controlled manipulation of DNA. Unfortunately, the K12 endonuclease turned out to be
perverse in its properties. While the enzyme does
bind to a defined recognition sequence, cleavage
occurs at a ‘random’ site several kilobases away
(Yuan et al. 1980). The much sought-after breakthrough finally came in 1970 with the discovery
in Haemophilus influenzae (Kelly & Smith 1970,
Smith & Wilcox 1970) of an enzyme that behaves
more simply. That is, the enzyme recognizes a particular target sequence in a duplex DNA molecule
and breaks the polynucleotide chain within that
sequence to give rise to discrete fragments of defined
length and sequence.
The presence of restriction and modification
systems is a double-edged sword. On the one hand,
they provide a rich source of useful enzymes for
DNA manipulation. On the other, these systems can
significantly affect the recovery of recombinant DNA
in cloning hosts. For this reason, some knowledge of
restriction and modification is essential.
Host-controlled restriction
and modification
Restriction systems allow bacteria to monitor the
origin of incoming DNA and to destroy it if it is
recognized as foreign. Restriction endonucleases
recognize specific sequences in the incoming DNA
and cleave the DNA into fragments, either at specific
sites or more randomly. When the incoming DNA is
a bacteriophage genome, the effect is to reduce the
efficiency of plating, i.e. to reduce the number of
plaques formed in plating tests. The phenomena of
restriction and modification were well illustrated
and studied by the behaviour of phage λ on two
E. coli host strains.
If a stock preparation of phage λ, for example, is
made by growth upon E. coli strain C and this stock is
then titred upon E. coli C and E. coli K, the titres
observed on these two strains will differ by several
orders of magnitude, the titre on E. coli K being the
lower. The phage are said to be restricted by the second host strain (E. coli K). When those phage that do
result from the infection of E. coli K are now replated
on E. coli K they are no longer restricted; but if they
are first cycled through E. coli C they are once again
restricted when plated upon E. coli K (Fig. 3.1). Thus
the efficiency with which phage λ plates upon a
POGC03 9/11/2001 11:00 AM Page 27
Cutting and joining DNA molecules
E. coli K
EOP = 10–4
EOP = 1
EOP = 1
λ.K
EOP = 1
λ.C
E. coli C
Fig. 3.1 Host-controlled restriction and modification of phage
λ in E. coli strain K, analysed by efficiency of plating (EOP).
Phage propagated by growth on strains K or C (i.e. λ.K or λ.C)
have EOPs on the two strains, as indicated by arrows.
particular host strain depends upon the strain on
which it was last propagated. This non-heritable
change conferred upon the phage by the second
host strain (E. coli K) that allows it to be replated
on that strain without further restriction is called
modification.
The restricted phages adsorb to restrictive hosts
and inject their DNA normally. When the phage
are labelled with 32P, it is apparent that their DNA
is degraded soon after injection (Dussoix & Arber
1962) and the endonuclease that is primarily
responsible for this degradation is called a restriction
endonuclease or restriction enzyme (Lederberg &
Meselson 1964). The restrictive host must, of course,
protect its own DNA from the potentially lethal
effects of the restriction endonuclease and so its DNA
must be appropriately modified. Modification involves
methylation of certain bases at a very limited number of sequences within DNA, which constitute the
recognition sequences for the restriction endonuclease. This explains why phage that survive one cycle
of growth upon the restrictive host can subsequently
reinfect that host efficiently; their DNA has been
replicated in the presence of the modifying methylase and so it, like the host DNA, becomes methylated and protected from the restriction system.
Although phage infection has been chosen as our
example to illustrate restriction and modification,
these processes can occur whenever DNA is transferred from one bacterial strain to another.
Types of restriction and modification
(R-M) system
At least four different kinds of R-M system are
known: type I, type II, type III and type IIs. The
essential differences between them are summarized
in Table 3.1.
The type I systems were the first to be characterized and a typical example is that from E. coli K12.
The active enzyme consists of two restriction subunits, two modification (methylation) subunits and
one recognition subunit. These subunits are the
Table 3.1 Characteristics of the different types of endonucleases.
System
27
Key features
Type I
One enzyme with different subunits for recognition, cleavage and methylation. Recognizes and methylates a single
sequence but cleaves DNA up to 1000 bp away
Type II
Two different enzymes which both recognize the same target sequence, which is symmetrical. The two enzymes either
cleave or modify the recognition sequence
Type III
One enzyme with two different subunits, one for recognition and modification and one for cleavage. Recognizes and
methylates same sequence but cleaves 24–26 bp away
Type IIs
Two different enzymes but recognition sequence is asymmetric. Cleavage occurs on one side of recognition sequence
up to 20 bp away
POGC03 9/11/2001 11:00 AM Page 28
28
CHAPTER 3
Box 3.1 Restriction: from a phenomenon in
bacterial genetics to a biological revolution
In the two related phenomena of host-controlled
restriction and modification, a single cycle of phage
growth, in a particular host bacterium, alters the
host range of the progeny virus. The virus may fail
to plate efficiently on a second host; its host range
is restricted. This modification of the virus differs
fundamentally from mutation because it is imposed
by the host cell on which the virus has been grown
but it is not inherited; when the phage is grown
in some other host, the modification may be lost.
In the 1950s, restriction and modification were
recognized as common phenomena, affecting many
virulent and temperate (i.e. capable of forming
lysogens) phages and involving various bacterial
species (Luria 1953, Lederberg 1957).
Arber and Dussoix clarified the molecular basis
of the phenomena. They showed that restriction
of phage l is associated with rapid breakdown of
the phage DNA in the host bacterium. They also
showed that modification results from an alteration
of the phage DNA which renders the DNA insensitive
to restriction. They deduced that a single modified
strand in the DNA duplex is sufficient to prevent
restriction (Arber & Dussoix 1962, Dussoix &
Arber 1962). Subsequent experiments implicated
methylation of the DNA in the modification process
(Arber 1965).
Detailed genetic analysis, in the 1960s, of the
bacterial genes (in E. coli K and E. coli B) responsible
for restriction and modification supported the duality
of the two phenomena. Mutants of the bacteria could
be isolated that were both restriction-deficient and
modification-deficient (R−M−), or that were R−M+.
The failure to recover R+M− mutants was correctly
ascribed to the suicidal failure to confer protective
modification upon the host’s own DNA.
The biochemistry of restriction advanced with
the isolation of the restriction endonuclease from
E. coli K (Meselson & Yuan 1968). It was evident
that the restriction endonucleases from E. coli K
and E. coli B were important examples of proteins
that recognize specific structures in DNA, but the
properties of these type I enzymes as they are now
known, were complex. Although the recognition
sites in the phage could be mapped genetically
(Murray et al. 1973a), determined efforts to define
the DNA sequences cleaved were unsuccessful
(Eskin & Linn 1972, Murray et al. 1973b).
The breakthrough came with Hamilton Smith’s
discovery of a restriction endonuclease from
Haemophilus influenzae strain Rd (Smith & Wilcox
1970) and the elucidation of the nucleotide
sequence at its cleavage sites in phage T7 DNA
(Kelly & Smith 1970). This enzyme is now known as
HindII. The choice of T7 DNA as the substrate for
cleavage was a good one, because the bacterium
also contains another type II restriction enzyme,
HindIII, in abundance. Fortunately, HindIII does
not cleave T7 DNA, and so any contaminating
HindIII in the HindII preparation could not be
problematical (Old et al. 1975). Shortly after the
discovery of HindII, several other type II restriction
endonucleases were isolated and characterized.
EcoRI was foremost among these (Hedgepeth
et al. 1972). They were rapidly exploited in the first
recombinant DNA experiments.
G A A T T C
C T T A A G
By the mid-1960s, restriction and modification
had been recognized as important and interesting
phenomena within the field of bacterial genetics
(see, for example, Hayes 1968), but who could
have foreseen the astonishing impact of restriction
enzymes upon biology?
POGC03 9/11/2001 11:00 AM Page 29
Cutting and joining DNA molecules
products of the hsdR, hsdM and hsdS genes. The
methylation and cutting reactions both require ATP
and S-adenosylmethionine as cofactors. The recognition sequences are quite long with no recognizable
features such as symmetry. The enzyme also cuts
unmodified DNA at some distance from the recognition sequence. However, because the methylation
reaction is performed by the same enzyme which
mediates cleavage, the target DNA may be modified
before it is cut. These features mean that type I systems are of little value for gene manipulation (see
also Box 3.1). However, their presence in E. coli
strains can affect recovery of recombinants (see
p. 33). Type III enzymes have symmetrical recognition sequences but otherwise resemble type I
systems and are of little value.
Most of the useful R-M systems are of type II. They
have a number of advantages over type I and III
systems. First, restriction and modification are mediated by separate enzymes so it is possible to cleave
DNA in the absence of modification. Secondly, the
restriction activities do not require cofactors such as
ATP or S-adenosylmethionine, making them easier
to use. Most important of all, type II enzymes recognize a defined, usually symmetrical, sequence and
cut within it. Many of them also make a staggered
break in the DNA and the usefulness of this will
become apparent. Although type IIs systems have
similar cofactors and macromolecular structure to
those of type II systems, the fact that restriction
occurs at a distance from the recognition site limits
their usefulness.
The classification of R-M systems into types I to III
is convenient but may require modification as new
discoveries are made. For example, the Eco571 system comprises a single polypeptide which has both
29
restriction and modification activities (Petrusyte
et al. 1988). Other restriction systems are known
which fall outside the above classification. Examples
include the mcr and mrr systems (see p. 34) and
homing endonucleases. The latter are doublestranded DNases derived from introns or inteins
(Belfort & Roberts 1997). They have large, asymmetric recognition sequences and, unlike standard
restriction endonucleases, tolerate some sequence
degeneracy within their recognition sequence.
Nomenclature
The discovery of a large number of restriction and
modification systems called for a uniform system of
nomenclature. A suitable system was proposed by
Smith and Nathans (1973) and a simplified version
of this is in use today. The key features are:
• The species name of the host organism is identified by the first letter of the genus name and the first
two letters of the specific epithet to generate a threeletter abbreviation. This abbreviation is always
written in italics.
• Where a particular strain has been the source
then this is identified.
• When a particular host strain has several different
R-M systems, these are identified by roman numerals.
Some examples are given in Table 3.2.
Homing endonucleases are named in a similar
fashion except that intron-encoded endonucleases
are given the prefix ‘I-’ (e.g. I-CeuI) and intein
endonucleases have the prefix ‘PI-’ (e.g. Pl-PspI).
Where it is necessary to distinguish between the
restriction and methylating activities, they are given
the prefixes ‘R’ and ‘M’, respectively, e.g. R.SmaI and
M.SmaI.
Table 3.2 Examples of restriction endonuclease nomenclature.
Enzyme
SmaI
HaeIII
HindII
HindIII
BamHI
Enzyme source
Recognition sequence
Serratia marcescens, 1st enzyme
Haemophilus aegyptius, 3rd enzyme
Haemophilus influenzae, strain d, 2nd enzyme
Haemophilus influenzae, strain d, 3rd enzyme
Bacillus amyloliquefaciens, strain H, 1st enzyme
CCCGGG
GGCC
GTPyPuAC
AAGCTT
GGATCC
POGC03 9/11/2001 11:00 AM Page 30
30
CHAPTER 3
These DNA fragments can associate by hydrogen
bonding between overlapping 5′ termini, or the fragments can circularize by intramolecular reaction
(Fig. 3.2). For this reason the fragments are said to
have sticky or cohesive ends. In principle, DNA fragments from diverse sources can be joined by means
of the cohesive ends and, as we shall see later, the
nicks in the molecules can be sealed to form an intact
artificially recombinant DNA molecule.
Not all type II enzymes cleave their target sites like
EcoRI. Some, such as PstI (CTGCA/G), produce fragments bearing 3′ overhangs, while others, such as
SmaI (CCC/GGG), produce blunt or flush ends.
To date, over 10 000 microbes from around the
world have been screened for restriction enzymes.
From these, over 3000 enzymes have been found
representing approximately 200 different sequence
specificities. Some representative examples are shown
in Table 3.3. For a comprehensive database of
information on restriction endonucleases and their
associated methylases, including cleavage sites,
commercial availability and literature references,
the reader should consult the website maintained by
New England Biolabs (www.rebase.neb.com).
Occasionally enzymes with novel DNA sequence
specificities are found but most prove to have the
same specificity as enzymes already known. Restriction enzymes with the same sequence specificity and
cut site are known as isoschizomers. Enzymes that
recognize the same sequence but cleave at different
points, for example SmaI (CCC/GGG) and XmaI
C/CCGGG), are sometimes known as neoschizomers.
Under extreme conditions, such as elevated pH
or low ionic strength, restriction endonucleases are
Recognition sequences
Most, but not all, type II restriction endonucleases
recognize and cleave DNA within particular sequences
of four to eight nucleotides which have a twofold
axis of rotational symmetry. Such sequences are often
referred to as palindromes because of their similarity
to words that read the same backwards as forwards.
For example, the restriction and modification enzymes
R.EcoRI and M.EcoRI recognize the sequence:
5′-GAA T T C-3′
3′-C T T AAG-5′
Axis of symmetry
The position at which the restricting enzyme cuts is
usually shown by the symbol ‘/’ and the nucleotides
methylated by the modification enzyme are usually
marked with an asterisk. For EcoRI these would be
represented thus:
5′-G/AA* T T C-3′
3′-C T T A*A/G-5′
For convenience it is usual practice to simplify the
description of recognition sequences by showing
only one strand of DNA, that which runs in the 5′ to
3′ direction. Thus the EcoRI recognition sequence
would be shown as G/AATTC.
From the information shown above we can see
that EcoRI makes single-stranded breaks four bases
apart in the opposite strands of its target sequence so
generating fragments with protruding 5′ termini:
5′-G
3′-CTTAA-5′
5′-AATTC-3′
G-5′
Intermolecular associatIon
5‘
Intramolecular association
AATT
TTAA
5‘
5‘
5‘
TTAA
AATT
AATT
AATT
AA
T TA T
T
A
AATT
TTAA
5‘
Base-pairing
5‘
Base-pairing
5‘
AATT
TTAA
5‘
Fig. 3.2 Cohesive ends of DNA fragments produced
by digestion with EcoRI.
POGC03 9/11/2001 11:00 AM Page 31
Cutting and joining DNA molecules
Table 3.3 Some restriction endonucleases and their
recognition sites.
Enzyme
Recognition sequence
4-Base cutters
MboI, DpnI, Sau3AI
MspI, HpaII
AluI
HaeIII
TaiI
/GATC
C /CGG
AG /CT
GG /CC
ACGT/
6-Base cutters
BglII
ClaI
PvuII
PvuI
KpnI
A/GATCT
AT/CGAT
CAG/CTG
CGAT/CG
GGTAC /C
8-Base cutters
NotI
Sbf I
GC /GGCCGC
CCTGCA/GG
capable of cleaving sequences which are similar but
not identical to their defined recognition sequence.
This altered specificity is known as star activity. The
most common types of altered activity are acceptance of base substitutions and truncation of the
number of bases in the recognition sequence. For
example, EcoRI* (EcoRI star activity) cleaves the
sequence N/AATTN, where N is any base, whereas
EcoRI cleaves the sequence GAATTC.
Number and size of restriction fragments
The number and size of the fragments generated
by a restriction enzyme depend on the frequency of
31
occurrence of the target site in the DNA to be cut.
Assuming a DNA molecule with a 50% G+C content
and a random distribution of the four bases, a fourbase recognition site occurs every 44 (256) bp.
Similarly, a six-base recognition site occurs every 46
(4096) bp and an eight-base recognition sequence
every 48 (65 536) bp. In practice, there is not a random distribution of the four bases and many organisms can be AT- or GC-rich, e.g. the nuclear genome
of mammals is 40% G+C and the dinucleotide CG is
fivefold less common than statistically expected.
Similarly, CCG and CGG are the rarest trinucleotides
in most A+T-rich bacterial genomes and CTAG is the
rarest tetranucleotide in G+C-rich bacterial genomes.
Thus different restriction endonucleases with six-base
recognition sites can produce average fragment sizes
significantly different from the expected 4096 bp
(Table 3.4).
Certain restriction endonucleases show preferential cleavage of some sites in the same DNA
molecule. For example, phage λ DNA has five sites
for EcoRI but the different sites are cleaved nonrandomly (Thomas & Davis 1975). The site nearest
the right terminus is cleaved 10 times faster than the
sites in the middle of the molecule. There are four
sites for SacII in λ DNA but the three sites in the
centre of the molecule are cleaved 50 times faster
than the remaining site. There is a group of three
restriction enzymes which show an even more
dramatic site preference. These are NarI, NaeI and
SacII and they require simultaneous interaction with
two copies of their recognition sequence before they
will cleave DNA (Kruger 1988, Conrad & Topal,
1989). Thus NarI will rapidly cleave two of the four
recognition sites on plasmid pBR322 DNA but will
seldom cleave the remaining two sites.
Table 3.4 Average fragment size (bp) produced by different enzymes with DNA from different sources.
Enzyme
ApaI
AvrII
BamHI
DraI
SpeI
Target
Arabidopsis
Nematode
Drosophila
E. coli
Human
GGGCCC
CCTAGG
GGATCC
TTTAAA
ACTAGT
25 000
15 000
6 000
2 000
8 000
40 000
20 000
9 000
1 000
8 000
6 000
20 000
4 000
1 000
9 000
15 000
150 000
5 000
2 000
60 000
2 000
8 000
5 000
2 000
10 000
POGC03 9/11/2001 11:00 AM Page 32
32
CHAPTER 3
Variations on cutting and
joining DNA molecules
In order to join two fragments of DNA together, it is
not essential that they are produced by the same
restriction endonuclease. Many different restriction
endonucleases produce compatible cohesive ends.
For example, AgeI (A/CCGGT) and AvaI (C/CCGGG)
produce molecules with identical 5′ overhangs and
so can be ligated together. There are many other
examples of compatible cohesive ends. What is
more, if the cohesive ends were produced by six-base
cutters, the ligation products are often recleavable
by four-base cutters. Thus, in the example cited
above, the hybrid site ACCGGG can be cleaved by
HpaII (C/CGG), NciI (CC/GGG) or ScrFI (CC/NGG).
New restriction sites can be generated by filling in
the overhangs generated by restriction endonucleases and ligating the products together. Figure 3.3
shows that after filling in the cohesive ends produced
by EcoRI, ligation produces restriction sites recognized by four other enzymes. Many other examples
of creating new target sites by filling and ligation
are known.
There are also many examples of combinations of
blunt-end restriction endonucleases that produce
recleavable ligation products. For example, when
molecules generated by cleavage with AluI (AG/CT)
are joined to ones produced by EcoRV (GAT/ATC),
some of the ligation sites will have the sequence
GATCT and others will have the sequence AGATC.
Both can be cleaved by MboI (GATC).
GAATTC
CTTAAG
EcoRI
AATTC
G
CTTAA
G
DNA polymerase
GAATT
AATTC
CTTAA
TTAAG
DNA ligase
GAATTAATTC
CTTAATTAAG
XmnI
AseI or
MseI
Tsp509 I
GAATT AATTC
GAAT
CTTAA TTAAG
CTTAAT
G
AseI AT/TAAT Tsp509 I
/AATT
MseI T/TAA
GAANN/NNTTC
XmnI
TAAG
AATTC
CTTAA
G
(Identical to EcoRI cleavage)
Recognition Sequences
TAATTC
Fig. 3.3 The generation of three new
restriction sites after filling in the
overhangs produced by endonuclease
EcoRI and ligating the products
together. Note that there are
two target sites, 4 bp apart, in the
reconstituted molecule
for endonuclease Tsp509I.
POGC03 9/11/2001 11:00 AM Page 33
Cutting and joining DNA molecules
A methyltransferase, M.SssI, that methylates the
dinucleotide CpG (Nur et al. 1985) has been isolated
from Spiroplasma. This enzyme can be used to modify
in vitro restriction endonuclease target sites which
contain the CG sequence. Some of the target
sequences modified in this way will be resistant to
endonuclease cleavage, while others will remain
sensitive. For example, if the sequence CCGG is
modified with SssI, it will be resistant to HpaII but
sensitive to MspI. Since 90% of the methyl groups in
the genomic DNA of many animals, including vertebrates and echinoderms, occur as 5-methylcytosine
in the sequence CG, M.Sss can be used to imprint
DNA from other sources with a vertebrate pattern.
The Dam and Dcm methylases of E. coli
Most laboratory strains of E. coli contain three sitespecific DNA methylases. The methylase encoded
by the dam gene transfers a methyl group from
S-adenosylmethionine to the N6 position of the
adenine residue in the sequence GATC. The methylase encoded by the dcm gene (the Dcm methylase,
previously called the Mec methylase) modifies the
internal cytosine residues in the sequences CCAGG
and CCTGG at the C5 position (Marinus et al. 1983).
In DNA in which the GC content is 50%, the sites
for these two methylases occur, on average, every
256–512 bp. The third methylase is the enzyme
M.EcoKI but the sites for this enzyme are much rarer
and occur about once every 8 kb.
These enzymes are of interest for two reasons.
First, some or all of the sites for a restriction endonuclease may be resistant to cleavage when
isolated from strains expressing the Dcm or Dam
methylases. This occurs when a particular base in
the recognition site of a restriction endonuclease
is methylated. The relevant base may be methylated
by one of the E. coli methylases if the methylase
recognition site overlaps the endonuclease recognition site. For example, DNA isolated from Dam+
E. coli is completely resistant to cleavage by MboI, but
not Sau3AI, both of which recognize the sequence
GATC. Similarly, DNA from a Dcm+ strain will be
cleaved by BstNI but not by EcoRII, even though
both recognize the sequence CCATGG. It is worth
noting that most cloning strains of E. coli are Dam+
Dcm+ but double mutants are available (Marinus
et al. 1983).
33
The second reason these methylases are of interest is that the modification state of plasmid DNA can
affect the frequency of transformation in special situations. Transformation efficiency will be reduced when
Dam-modified plasmid DNA is introduced into Dam−
E. coli or Dam- or Dcm-modified DNA is introduced into
other species (Russell & Zinder 1987). When DNA is
to be moved from E. coli to another species it is best to
use a strain lacking the Dam and Dcm methylases.
As will be seen later, it is difficult to stably clone
DNA that contains short, direct repetitive sequences.
Deletion of the repeat units occurs quickly, even
when the host strain is deficient in recombination.
However, the deletion mechanism appears to involve
Dam methylation, for it does not occur in dam
mutants (Troester et al. 2000).
The importance of eliminating
restriction systems in E. coli strains used
as hosts for recombinant molecules
If foreign DNA is introduced into an E. coli host it
may be attacked by restriction systems active in the
host cell. An important feature of these systems is
that the fate of the incoming DNA in the restrictive
host depends not only on the sequence of the DNA
but also upon its history: the DNA sequence may or
may not be restricted, depending upon its source
immediately prior to transforming the E. coli host
strain. As we have seen, post-replication modifications
of the DNA, usually in the form of methylation of particular adenine or cytosine residues in the target sequence, protect against cognate restriction systems but
not, in general, against different restriction systems.
Because restriction provides a natural defence
against invasion by foreign DNA, it is usual to
employ a K restriction-deficient E. coli K12 strain
as a host in transformation with newly created
recombinant molecules. Thus where, for example,
mammalian DNA has been ligated into a plasmid
vector, transformation of the EcoK restrictiondeficient host eliminates the possibility that the
incoming sequence will be restricted, even if the
mammalian sequence contains an unmodified EcoK
target site. If the host happens to be EcoK restrictiondeficient but EcoK modification-proficient, propagation
on the host will confer modification methylation and
hence allow subsequent propagation of the recombinant in EcoK restriction-proficient strains, if desired.
POGC03 9/11/2001 11:00 AM Page 34
34
CHAPTER 3
mcrC
mcrB
hsdS
hsdM
hsdR
14 kb
Whereas the EcoKI restriction system, encoded by
the hsdRMS genes, cleaves DNA that is not protected
by methylation at the target site, the McrA, McrBC
and Mrr endonucleases cut DNA that is methylated
at specific positions. All three endonucleases restrict
DNA modified by CpC methylase (M.SssI) and the
Mrr endonuclease will attack DNA with methyladenine in specific sequences. The significance of these
restriction enzymes is that DNA from many bacteria,
and from all plants and higher animals, is extensively
methylated and its recovery in cloning experiments
will be greatly reduced if the restriction activity is
not eliminated. There is no problem with DNA from
Saccharomyces cerevisiae or Drosophila melanogaster
since there is little methylation of their DNA.
All the restriction systems in E. coli are clustered
together in an ‘immigration control region’ about
14 kb in length (Fig. 3.4). Some strains carry mutations in one of the genes. For example, strains DH1
and DH5 have a mutation in the hsdR gene and so
are defective for the EcoKI endonuclease but still
mediate the EcoKI modification of DNA. Strain DP50
has a mutation in the hsdS gene and so lacks both the
EcoKI restriction and modification activities. Other
strains, such as E. coli C and the widely used cloning
strain HB101, have a deletion of the entire mcrC–
mrr region and hence lack all restriction activities.
mrr
Fig. 3.4 The immigration control region of
E. coli strain K12.
where subsequent ligation is achieved, the resulting
product may contain small deletions. The message is
clear: cheap restriction enzymes are in reality poor
value for money!
A typical QC procedure is as follows. DNA fragments are produced by an excessive overdigestion of
substrate DNA with each restriction endonuclease.
These fragments are then ligated and recut with the
same restriction endonuclease. Ligation can occur
only if the 3′ and 5′ termini are left intact, and only
those molecules with a perfectly restored recognition site can be recleaved. A normal banding pattern
after cleavage indicates that both the 3′ and 5′ termini
are intact and the enzyme preparation is free of detectable exonucleases and phosphatases (Fig. 3.5).
The importance of enzyme quality
Restriction enzymes are available from many different commercial sources. In choosing a source of
enzyme, it is important to consider the quality of the
enzyme supplied. High-quality enzymes are purified
extensively to remove contaminating exonucleases
and endonucleases and tests for the absence of such
contaminants form part of routine quality control
(QC) on the finished product (see below). The absence
of exonucleases is particularly important. If they are
present, they can nibble away the overhangs of
cohesive ends, thereby eliminating or reducing the
production of subsequent recombinants. Contaminating phosphatases can remove the terminal phosphate residues, thereby preventing ligation. Even
Fig. 3.5 Quality control of the enzyme PstI. DNA was
overdigested with the endonuclease and the fragments were
ligated together and then recut. Note that the two digests give
an identical banding pattern upon agarose gel electrophoresis.
POGC03 9/11/2001 11:00 AM Page 35
Cutting and joining DNA molecules
35
E. coli. The transformants are plated on media containing the β-galactosidase substrate Xgal. If the
lacZ gene remains intact after digestion and ligation,
it will give rise to a blue colony. If any degradation
of the cut ends occurred, then a white colony will
be produced (Box 3.2).
An additional QC test is the blue/white screening
assay. In this, the starting material is a plasmid
carrying the E. coli lacZ gene in which there is a single recognition site for the enzyme under test. The
plasmid is overdigested with the restriction enzyme,
religated and transformed into a LacZ − strain of
Box 3.2 a-Complementation of b-galactosidase and the use of Xgal
The activity of the enzyme b-galactosidase is easily
monitored by including in the growth medium the
chromogenic substrate 5-bromo-4-chloro-3-indolylb-D-galactoside (Xgal). This compound is colourless
but on cleavage releases a blue indolyl derivative.
On solid medium, colonies that are expressing active
b-galactosidase are blue in colour while those
without the activity are white in colour. This is often
referred to as blue/white screening. Since Xgal is
not an inducer of b-galactosidase, the non-substrate
(gratuitous) inducer isopropyl-b-D-thiogalactoside
(IPTG) is also added to the medium.
CH2OH
O
HO
O
Cl
5-Bromo-4-chloro-3-indolylβ-D-galactoside (Xgal)
Br
OH
H
N
H
OH
β-Galactosidase
O
Cl
OH
Br
O
O
Cl
Br
Br
Air
N
H
5-Bromo-4-chloroindoxyl
The phenomenon of a-complementation of
b-galactosidase is widely used in molecular
genetics. The starting-point for a-complementation
is the M15 mutant of E. coli. This has a deletion
of residues 11– 41 in the lacZ gene and shows no
b-galactosidase activity. Enzyme activity can be
restored to the mutant enzyme in vitro by adding
a cyanogen bromide peptide derived from amino
acid residues 3–92 (Langley et al. 1975, Langley
& Zabin 1976). Complementation can also be
shown in vivo. If a plasmid carrying the N-terminal
fragment of the lacZ gene encompassing the missing
region is introduced into the M15 mutant, then
b-galactosidase is produced, as demonstrated by the
production of a blue colour on medium containing
N
H
N
H
5,5‘-Dibromo-4,4‘-dichloroindigo
Xgal. In practice, the plasmid usually carries the
lacI gene and the first 146 codons of the lacZ gene,
because in the early days of genetic engineering
this was a convenient fragment of DNA to
manipulate.
Since wild-type b-galactosidase has 1021 amino
acids, it is encoded by a gene 3.1 kb in length.
While a gene of this length is easily manipulated
in vitro, there are practical disadvantages to using
the whole gene. As will be seen later, it is preferable
to keep cloning vectors and their inserts as small as
possible. The phenomenon of a-complementation
allows genetic engineers to take advantage of the
lac system without having to have the entire Z gene
on the vector.
POGC03 9/11/2001 11:00 AM Page 36
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CHAPTER 3
Joining DNA molecules
Having described the methods available for cutting
DNA molecules, we must consider the ways in which
DNA fragments can be joined to create artificially
recombinant molecules. There are currently three
methods for joining DNA fragments in vitro. The first
of these capitalizes on the ability of DNA ligase
to join covalently the annealed cohesive ends produced by certain restriction enzymes. The second
depends upon the ability of DNA ligase from phage
T4-infected E. coli to catalyse the formation of phosphodiester bonds between blunt-ended fragments.
The third utilizes the enzyme terminal deoxynucleotidyltransferase to synthesize homopolymeric 3′
single-stranded tails at the ends of fragments. We
can now look at these three methods a little more
deeply.
DNA ligase
E. coli and phage T4 encode an enzyme, DNA ligase,
which seals single-stranded nicks between adjacent
P
5’
B
B
B
B
B
B
P
P
3’
P
OH
P
P
B
B
P
P
B
B
P
P
B
B
P
nucleotides in a duplex DNA chain (Olivera et al.
1968, Gumport & Lehman 1971). Although the
reactions catalysed by the enzymes of E. coli and
T4-infected E. coli are very similar, they differ in their
cofactor requirements. The T4 enzyme requires
ATP, while the E. coli enzyme requires NAD+. In
each case the cofactor is split and forms an enzyme –
AMP complex. The complex binds to the nick, which
must expose a 5′ phosphate and 3′ OH group, and
makes a covalent bond in the phosphodiester chain,
as shown in Fig. 3.6.
When termini created by a restriction endonuclease that creates cohesive ends associate, the joint
has nicks a few base pairs apart in opposite strands.
DNA ligase can then repair these nicks to form an
intact duplex. This reaction, performed in vitro
with purified DNA ligase, is fundamental to many
gene-manipulation procedures, such as that shown
in Fig. 3.7.
The optimum temperature for ligation of nicked
DNA is 37°C, but at this temperature the hydrogenbonded join between the sticky ends is unstable.
EcoRI-generated termini associate through only
3’
ATP
B
B
P
PPi
T4 DNA ligase
5’
Enzyme – AMP
A
Enzyme
P
P
5’
P
B
B
B
B
P
3’
P
OH
B
B
P
P
B
B
P
P
B
B
P
P
B
B
P
3’
E.coli DNA ligase
NMN
NAD+
B
B
P
5’
AMP
P
5’
B
B
3’
P
B
B
P
P
B
B
P
P
B
B
P
P
B
B
P
P
B
B
P
3’
B
B
P
5’
Fig. 3.6 Action of DNA ligase. An enzyme
– AMP complex binds to a nick bearing 3′
OH and 5′ P groups. The AMP reacts with
the phosphate group. Attack by the 3′
OH group on this moiety generates a new
phosphodiester bond, which seals the nick.
POGC03 9/11/2001 11:00 AM Page 37
Cutting and joining DNA molecules
37
Origin of
replication
Cleavage site
Foreign DNA
Plasmid
vector
Tc
resistance
Cleavage
site
Restriction
endonuclease
Restriction
endonuclease
AATT
AATT
TTA
A
TTAA
TTAA
A
A
TT
Annealing
AA
T
TT
T
A
A
TT
AA
A
A
TT
Recombinant
plasmid
Recombinant
plasmid
Chromosome
DNA ligase
seals nicks
Transformed bacterial cell
Fig. 3.7 Use of DNA ligase to create a
covalent DNA recombinant joined
through association of termini
generated by EcoRI.
Transformation, Tc selection
four AT base pairs and these are not sufficient to
resist thermal disruption at such a high temperature. The optimum temperature for ligating the
cohesive termini is therefore a compromise between
the rate of enzyme action and association of the
termini, and has been found experimentally to be
in the range 4–15°C (Dugaiczyk et al. 1975, Ferretti
& Sgaramella 1981).
The ligation reaction can be performed so as to
favour the formation of recombinants. First, the
population of recombinants can be increased by
performing the reaction at a high DNA concentration; in dilute solutions circularization of linear fragments is relatively favoured because of the reduced
frequency of intermolecular reactions. Secondly, by
treating linearized plasmid vector DNA with alkaline phosphatase to remove 5′-terminal phosphate
groups, both recircularization and plasmid dimer
formation are prevented (Fig. 3.8). In this case,
circularization of the vector can occur only by insertion of non-phosphatase-treated foreign DNA which
provides one 5′-terminal phosphate at each join.
One nick at each join remains unligated, but, after
transformation of host bacteria, cellular repair
mechanisms reconstitute the intact duplex.
Joining DNA fragments with cohesive ends by
DNA ligase is a relatively efficient process which has
been used extensively to create artificial recombinants. A modification of this procedure depends upon
the ability of T4 DNA ligase to join blunt-ended DNA
molecules (Sgaramella 1972). The E. coli DNA ligase
will not catalyse blunt ligation except under special
reaction conditions of macromolecular crowding
(Zimmerman & Pheiffer 1983). Blunt ligation is most
usefully applied to joining blunt-ended fragments
via linker molecules; in an early example of this,
Scheller et al. (1977) synthesized self-complementary
decameric oligonucleotides, which contain sites for
POGC03 9/11/2001 11:00 AM Page 38
38
CHAPTER 3
P
OH
P
Ligase
OH
+ dimers, etc.
Alkaline
phosphatase
2Pi
OH OH
Ligase
OHOH
HO
P
P
HO
No reaction
Ligase
Foreign DNA
fragment
Nick
Transformation
Nick
Host repairs one nick at each
join
one or more restriction endonucleases. One such
molecule is shown in Fig. 3.9. The molecule can be
ligated to both ends of the foreign DNA to be cloned,
and then treated with restriction endonuclease to
produce a sticky-ended fragment, which can be
incorporated into a vector molecule that has been
cut with the same restriction endonuclease. Insertion by means of the linker creates restrictionenzyme target sites at each end of the foreign DNA
and so enables the foreign DNA to be excised and
recovered after cloning and amplification in the host
bacterium.
Adaptors
It may be the case that the restriction enzyme used to
generate the cohesive ends in the linker will also cut
the foreign DNA at internal sites. In this situation,
Fig. 3.8 Application of alkaline
phosphatase treatment to prevent
recircularization of vector plasmid
without insertion of foreign DNA.
the foreign DNA will be cloned as two or more subfragments. One solution to this problem is to choose
another restriction enzyme, but there may not be a
suitable choice if the foreign DNA is large and has
sites for several restriction enzymes. Another solution is to methylate internal restriction sites with the
appropriate modification methylase. An example of
this is described in Fig. 6.2. Alternatively, a general
solution to the problem is provided by chemically
synthesized adaptor molecules which have a preformed cohesive end (Wu et al. 1978). Consider a
blunt-ended foreign DNA containing an internal
BamHI site (Fig. 3.10), which is to be cloned in a
BamHl-cut vector. The Bam adaptor molecule has
one blunt end bearing a 5′ phosphate group and a
Bam cohesive end which is not phosphorylated. The
adaptor can be ligated to the foreign DNA ends. The
foreign DNA plus added adaptors is then phosphory-
POGC03 9/11/2001 11:00 AM Page 39
EcoRI
Decameric
linker
molecule
C C G A A T T C G G 3’
G G C T T A A G C C 5’
T4 DNA
ligase
+
Linker
molecules
Foreign
DNA
EcoRI
DNA
ligase
Vector
Fig. 3.9 A decameric linker molecule
containing an EcoRI target site is joined
by T4 DNA ligase to both ends of flushended foreign DNA. Cohesive ends are
then generated by EcoRI. This DNA can
then be incorporated into a vector that
has been treated with the same
restriction endonuclease.
5’–HO –GATCCCCGGG –OH
3’–
HO –GGGCCC –P
Adaptor molecule
P
HO
HO
HO
OH
+
P
HO
HO
OH
P
OH
P
T4 DNA
ligase,
ATP
HO
HO
OH
OH
Polynucleotide
kinase, ATP
OH
P
Vector
OH P
Fig. 3.10 Use of a BamHI adaptor
molecule. A synthetic adaptor molecule
is ligated to the foreign DNA. The
adaptor is used in the 5′-hydroxyl form
to prevent self-polymerization. The
foreign DNA plus ligated adaptors is
phosphorylated at the 5′-termini and
ligated into the vector previously cut
with BamHI.
P
HO
OH
P
POGC03 9/11/2001 11:00 AM Page 40
40
CHAPTER 3
lated at the 5′ termini and ligated into the BamHI site
of the vector. If the foreign DNA were to be recovered
from the recombinant with BamHI, it would be
obtained in two fragments. However, the adaptor is
designed to contain two other restriction sites (SmaI,
HpaII), which may enable the foreign DNA to be
recovered intact.
Note that the only difference between an adaptor
and a linker is that the former has cohesive ends and
the latter has blunt ends. A wide range of adaptors
are available commercially.
Homopolymer tailing
A general method for joining DNA molecules makes
use of the annealing of complementary homopolymer
sequences. Thus, by adding oligo(dA) sequences to
the 3′ ends of one population of DNA molecules and
oligo(dT) blocks to the 3′ ends of another population,
the two types of molecule can anneal to form mixed
dimeric circles (Fig. 3.11).
An enzyme purified from calf thymus, terminal
deoxynucleotidyltransferase, provides the means
5’
3’
3’
5’
5’
3’
3’
5’
λ-exonuclease
λ-exonuclease
5’
3’
3’
5’
5’
3’
Terminal
transferase
+dATP
3’A(A)nA
5’
5’
by which the homopolymeric extensions can be
synthesized, for if presented with a single deoxynucleotide triphosphate it will repeatedly add nucleotides to the 3′ OH termini of a population of DNA
molecules (Chang & Bollum 1971). DNA with
exposed 3′ OH groups, such as arise from pretreatment with phage λ exonuclease or restriction with
an enzyme such as PstI, is a very good substrate
for the transferase. However, conditions have been
found in which the enzyme will extend even the
shielded 3′ OH of 5′ cohesive termini generated by
EcoRI (Roychoudhury et al. 1976, Humphries et al.
1978).
The terminal transferase reactions have been
characterized in detail with regard to their use in
gene manipulation (Deng & Wu 1981, Michelson
& Orkin 1982). Typically, 10– 40 homopolymeric
residues are added to each end.
One of the earliest examples of the construction of
recombinant molecules, the insertion of a piece of λ
DNA into SV40 viral DNA, made use of homopolymer tailing ( Jackson et al. 1972). In their experiments, the single-stranded gaps which remained
A(A)nA 3’
3’
5’
Terminal
transferase
+dATP
5’
3’T(T)nT
5’
T(T)nT 3’
Mix and
anneal
) nA
A(A ) nT
T(T
)T
T(T n A
)n
A(A
Fig. 3.11 Use of calf-thymus terminal
deoxynucleotidyltransferase to add
complementary homopolymer tails to
two DNA molecules.
POGC03 9/11/2001 11:00 AM Page 41
Cutting and joining DNA molecules
in the two strands at each join were repaired in
vitro with DNA polymerase and DNA ligase so as to
produce covalent-ly closed circular molecules. The
recombinants were then transfected into susceptible
mammalian cells (see Chapter 10). Subsequently,
the homopolymer method, using either dA.dT or
dG.dC homopolymers was used extensively to construct recombinant plasmids for cloning in E. coli. In
recent years, homopolymer tailing has been largely
replaced as a result of the availability of a much
wider range of restriction endonucleases and other
DNA-modifying enzymes. However, it is still important for cDNA cloning (see p. 95 et seq.).
Joining polymerase chain
reaction (PCR) products
Many of the strategies for cloning DNA fragments do
not work well with PCR products. The reason for this
is that the polymerases used in the PCR have a terminal transferase activity. For example, the Taq
polymerase adds a single 3′ A overhang to each end
of the PCR product. Thus PCR products cannot be
blunt-end-ligated unless the ends are first polished
(blunted). A DNA polymerase like Klenow can be
used to fill in the ends. Alternatively, Pfu DNA polymerase can be used to remove extended bases with
its 3′ to 5′ exonuclease activity. However, even
when the PCR fragments are polished, blunt-endligating them into a vector still may be very
inefficient. One solution to this problem is to use T/A
cloning (Mead et al. 1991). In this method, the PCR
fragment is ligated to a vector DNA molecule with a
single 3′ deoxythymidylate extension (Fig. 3.12).
41
Incorporation of extra sequence at the
5′′ end of a primer into amplified DNA
A PCR primer may be designed which, in addition
to the sequence required for hybridization with the
input DNA, includes an extra sequence at its 5′ end.
The extra sequence does not participate in the first
hybridization step – only the 3′ portion of the primer
hybridizes – but it subsequently becomes incorporated into the amplified DNA fragment (Fig. 3.13).
Because the extra sequence can be chosen at the
will of the experimenter, great flexibility is available
here.
A common application of this principle is the
incorporation of restriction sites at each end of the
amplified product. Figure 3.13 illustrates the addition of a HindIII site and an EcoRI site to the ends
of an amplified DNA fragment. In order to ensure
that the restriction sites are good substrates for
the restriction endonucleases, four nucleotides are
placed between the hexanucleotide restriction sites
and the extreme ends of the DNA. The incorporation
of these restriction sites provides one method for
cloning amplified DNA fragments (see below).
Joining DNA molecules without
DNA ligase
In all the cutting and joining reactions described
above, two separate protein components were
required: a site-specific endonuclease and a DNA ligase. Shuman (1994) has described a novel approach
to the synthesis of recombinant molecules in which
a single enzyme, vaccinia DNA topoisomerase, both
(a)
GG T G A NNNNNNNNNN – – – – –
C C A C T NNNNNNNNNN – – – – –
(b)
Fig. 3.12 Cleavage of a vector DNA
molecule to generate single thymidylate
overhangs. (a) The recognition
sequence and cleavage point for the
restriction endonuclease HphI.
(b) Sequences in the vector DNA which
result in desired overhangs after
cleavage with HphI.
HphI
GG T G A A C C CGGG T T CG A – – – – – – – – – – A G A A C C CGGG T T C A C C
C C A C T T GGGC CG A A GC T – – – – – – – – – – T C T T GGGC C C A A G T GG
HphI
HphI
GG T G A A C C CGGG T
C C A C T T GGGC C C
C C CGGG T T C A C C
T GGGC C C A A G T GG
POGC03 9/11/2001 11:00 AM Page 42
42
CHAPTER 3
Target region
5‘
3‘
3‘
5‘
Denature, anneal primers 1 and 2
5‘
3‘
3‘
Primer 1
5‘ G CG C A A G
C
TT
CTT
A AG CC GG 5‘
Primer 2
3‘
3‘
5‘
PCR amplification
with primers 1 and 2
HindIII
Target region
EcoRI
5‘ GCGC A A GC T T
G A A T T CGGC C
3‘ CGCG T T CG A A
C T T A A GC CGG
cleaves and rejoins DNA molecules. Placement of
the CCCTT cleavage motif for vaccinia topoisomerase near the end of a duplex DNA permits efficient
generation of a stable, highly recombinogenic protein
DNA adduct that can only religate to acceptor DNAs
that contain complementary single-strand extensions. Linear DNAs containing CCCTT cleavage sites
at both ends can be activated by topoisomerase and
inserted into a plasmid vector.
Heyman et al. (1999) have used the properties
of vaccinia topoisomerase to develop a ligase-free
Fig. 3.13 Incorporation of extra
sequence at the 5′ end of a primer.
Two primers have sequences designed
to hybridize at the ends of the target
region. Primer 1 has an extra sequence
near its 5′ end which forms a HindIII
site (AAGCTT), and primer 2 has an
extra sequence near its 5′ end which
forms an EcoRI (GAATTC) site. Each
primer has an additional 5′-terminal
sequence of four nucleotides so that
the hexanucleotide restriction sites are
placed within the extreme ends of
the amplified DNA, and so present
good substrates for endonuclease
cleavage.
technology for the covalent joining of DNA fragments
to plasmid vectors. Whereas joining molecules
with DNA ligase requires an overnight incubation,
topoisomerase-mediated ligation occurs in 5 min.
The method is particularly suited to cloning PCR
fragments. A linearized vector with single 3′ T
extensions is activated with the topoisomerase. On
addition of the PCR product with 3′ A overhangs,
ligation is very rapid. In addition, the high substrate
specificity of the enzyme means that there is a low
rate of formation of vectors without inserts.
POGC04 9/11/2001 10:59 AM Page 43
CHAPTER 4
Basic biology of plasmid
and phage vectors
Plasmid biology and
simple plasmid vectors
Plasmids are widely used as cloning vehicles but,
before discussing their use in this context, it is appropriate to review some of their basic properties.
Plasmids are replicons which are stably inherited in
an extrachromosomal state. Most plasmids exist as
double-stranded circular DNA molecules. If both
strands of DNA are intact circles the molecules are
described as covalently closed circles or CCC DNA
(Fig. 4.1). If only one strand is intact, then the
molecules are described as open circles or OC DNA.
When isolated from cells, covalently closed circles
often have a deficiency of turns in the double helix,
such that they have a supercoiled configuration.
Supercoiled
DNA
DNA gyrase
Endonuclease
Topoisomerase
Endonuclease
DNA ligase
Relaxed, covalently
closed circular DNA
The enzymatic interconversion of supercoiled, relaxed
CCC DNA* and OC DNA is shown in Fig. 4.1.
Because of their different structural configurations,
supercoiled and OC DNA separate upon electrophoresis in agarose gels (Fig. 4.2). Addition of an
intercalating agent, such as ethidium bromide, to
supercoiled DNA causes the plasmid to unwind. If
excess ethidium bromide is added, the plasmid will
rewind in the opposite direction (Fig. 4.3). Use of
this fact is made in the isolation of plasmid DNA
(see p. 48).
Not all plasmids exist as circular molecules.
Linear plasmids have been found in a variety of bacteria, e.g. Streptomyces sp. and Borrelia burgdorferi.
To prevent nuclease digestion, the ends of linear
plasmids need to be protected and two general
mechanisms have evolved. Either there are repeated
sequences ending in a terminal DNA hairpin loop
(Borrelia) or the ends are protected by covalent
attachment of a protein (Streptomyces). For more
details of linear plasmids the reader should consult
Hinnebusch and Tilly (1993).
Plasmids are widely distributed throughout the
prokaryotes, vary in size from less than 1 × 106
daltons to greater than 200 × 106, and are generally
dispensable. Some of the phenotypes which these
plasmids confer on their host cells are listed in
Table 4.1. Plasmids to which phenotypic traits have
not yet been ascribed are called cryptic plasmids.
Plasmids can be categorized into one of two major
type – conjugative or non-conjugative – depending
upon whether or not they carry a set of transfer
genes, called the tra genes, which promote bacterial
conjugation. Plasmids can also be categorized on the
basis of their being maintained as multiple copies per
Open, circular DNA
Fig. 4.1 The interconversion of supercoiled, relaxed
covalently closed circular DNA and open circular DNA.
* The reader should not be confused by the terms relaxed circle
and relaxed plasmid. Relaxed circles are CCC DNA that does not
have a supercoiled configuration. Relaxed plasmids are plasmids with multiple copies per cell.
POGC04 9/11/2001 10:59 AM Page 44
44
CHAPTER 4
A
cell (relaxed plasmids) or as a limited number of
copies per cell (stringent plasmids). Generally, conjugative plasmids are of relatively high molecular
weight and are present as one to three copies per
chromosome, whereas non-conjugative plasmids
are of low molecular weight and present as multiple
copies per chromosome (Table 4.2). An exception is
the conjugative plasmid R6K, which has a molecular weight of 25 × 106 daltons and is maintained as a
relaxed plasmid.
B
A
Direction of
migration
B
OC
L
C
SC
D
Fig. 4.2 Electrophoresis of DNA in agarose gels. The direction
of migration is indicated by the arrow. DNA bands have been
visualized by soaking the gel in a solution of ethidium bromide
(which complexes with DNA by intercalating between
stacked base pairs) and photographing the orange
fluorescence which results upon ultraviolet irradiation.
(A) Open circular (OC) and supercoiled (SC) forms of a plasmid
of 6.4 kb pairs. Note that the compact supercoils migrate
considerably faster than open circles (B). Linear plasmid
(L) DNA is produced by treatment of the preparation shown
in lane (A) with EcoRI, for which there is a single target site.
Under the conditions of electrophoresis employed here, the
linear form migrates just ahead of the open circular form.
E
Fig. 4.3 Effect of intercalation of ethidium bromide on
supercoiling of DNA. As the amount of intercalated ethidium
bromide increases, the double helix untwists, with the result
that the supercoiling decreases until the open form of the
circular molecule is produced. Further intercalation
introduces excess turns in the double helix, resulting in
supercoiling in the opposite sense (note the direction of
coiling at B and D). For clarity, only a single line represents
the double helix.
POGC04 9/11/2001 10:59 AM Page 45
Biology of plasmid and phage vectors
Table 4.1 Some phenotypic traits exhibited by
plasmid-carried genes.
Antibiotic resistance
Antibiotic production
Degradation of aromatic compounds
Haemolysin production
Sugar fermentation
Enterotoxin production
Heavy-metal resistance
Bacteriocin production
Induction of plant tumours
Hydrogen sulphide production
Host-controlled restriction and modification
Host range of plasmids
Plasmids encode only a few of the proteins required
for their own replication and in many cases encode
only one of them. All the other proteins required
for replication, e.g. DNA polymerases, DNA ligase,
helicases, etc., are provided by the host cell. Those
replication proteins that are plasmid-encoded are
located very close to the ori (origin of replication)
sequences at which they act. Thus, only a small
region surrounding the ori site is required for replication. Other parts of the plasmid can be deleted and
foreign sequences can be added to the plasmid and
replication will still occur. This feature of plasmids
has greatly simplified the construction of versatile
cloning vectors.
The host range of a plasmid is determined by its ori
region. Plasmids whose ori region is derived from
45
plasmid Col E1 have a restricted host range: they
only replicate in enteric bacteria, such as E. coli,
Salmonella, etc. Other promiscuous plasmids have
a broad host range and these include RP4 and
RSF1010. Plasmids of the RP4 type will replicate in
most Gram-negative bacteria, to which they are
readily transmitted by conjugation. Such promiscuous plasmids offer the potential of readily transferring cloned DNA molecules into a wide range of
genetic backgrounds. Plasmids like RSF1010 are
not conjugative but can be transformed into a wide
range of Gram-negative and Gram-positive bacteria,
where they are stably maintained. Many of the plasmids isolated from Staphylococcus aureus also have a
broad host range and can replicate in many other
Gram-positive bacteria. Plasmids with a broad host
range encode most, if not all, of the proteins required
for replication. They must also be able to express
these genes and thus their promoters and ribosome
binding sites must have evolved such that they can
be recognized in a diversity of bacterial families.
Plasmid copy number
The copy number of a plasmid is determined by
regulating the initiation of plasmid replication. Two
major mechanisms of control of initiation have been
recognized: regulation by antisense RNA and regulation by binding of essential proteins to iterons (for
review, see Del Solar et al. 1998). Most of the cloning
vectors in current use carry an ori region derived
from plasmid Col E1 and copy-number control is
mediated by antisense RNA. In this type of plasmid, the primer for DNA replication is a 555-base
Table 4.2 Properties of some conjugative and non-conjugative plasmids of Gram-negative organisms.
Plasmid
Size (MDa)
Conjugative
No. of plasmid copies/
chromosome equivalent
Phenotype
Col E1
RSF1030
clo DF13
R6K
F
RI
Ent P 307
4.2
5.6
6
25
62
62.5
65
No
No
No
Yes
Yes
Yes
Yes
10–15
20– 40
10
13–38
1–2
3–6
1–3
Col E1 production
Ampicillin resistance
Cloacin production
Ampicillin and streptomycin resistance
–
Multiple drug resistance
Enterotoxin production
POGC04 9/11/2001 10:59 AM Page 46
46
CHAPTER 4
Rop dimer
RNA I
ρ RNAI
ρrop
Origin
5‘
3‘
3‘
5‘
ρ RNAII
RNA II
Infrequently
Frequently
3‘
5‘
RNA II
RNA I
5‘
5‘
RNA II
RNase H
5‘
RNA II
3‘
5‘
3‘
5‘
RNA I-RNA II duplex
Pol I
RNA II primes
DNA synthesis
RNA II inactivated for
primer function
Replication
No replication
ribonucleotide molecule called RNA II, which forms
an RNA–DNA hybrid at the replication origin
(Tomizawa & Itoh 1982), RNA II can only act as
a primer if it is cleaved by RNase H to leave a free
3′ hydroxyl group. Unless RNA II is processed in this
way, it will not function as a primer and replication
will not ensue. Replication control is mediated by
another small (108-base) RNA molecule called RNA
I (Tomizawa & Itoh 1981), which is encoded by
the same region of DNA as RNA II but by the complementary strand. Thus RNA I and RNA II are
complementary to each other and can hybridize to
form a double-stranded RNA helix. The formation of
this duplex interferes with the processing of RNA II
by RNase H and hence replication does not ensue
Fig. 4.4 Regulation of replication of
Col E1-derived plasmids. RNA II must
be processed by RNase H before it can
prime replication. ‘Origin’ indicates
the transition point between the RNA
primer and DNA. Most of the time,
RNA I binds to RNA II and inhibits
the processing, thereby regulating the
copy number. ρRNAI and ρRNA II are
the promoters for RNA I and RNA II
transcription, respectively. RNA I is
coloured pale red and RNA II dark red.
The Rop protein dimer enhances the
initial pairing of RNA I and RNA II.
(Fig. 4.4). Since RNA I is encoded by the plasmid,
more of it will be synthesized when the copy number
of the plasmid is high. As the host cell grows and
divides, so the concentration of RNA I will fall and
the plasmid will begin to replicate again (Cesarini
et al. 1991, Eguchi et al. 1991).
In addition to RNA I, a plasmid-encoded protein
called Rop helps maintain the copy number (Cesarini
et al. 1982). This protein, which forms a dimer,
enhances the pairing between RNA I and RNA II so
that processing of the primer can be inhibited even
at relatively low concentrations of RNA I. Deletion of
the ROP gene (Twigg & Sherratt 1980) or mutations
in RNA I (Muesing et al. 1981) result in increased
copy numbers.
POGC04 9/11/2001 10:59 AM Page 47
Biology of plasmid and phage vectors
ori
bp
200
400
par
DnaA
600
800
inc
R1 R2
R3
1000
1200
repA
IR1
IR2
repA
–35 –10
mRNA
Fig. 4.5 The ori region of pSC101. R1, R2,
and R3 are the three iteron sequences
(CAAAGGTCTAGCAGCAGAATTTACAGA for R3) to which
RepA binds to handcuff two plasmids. RepA autoregulates its
own synthesis by binding to the inverted repeats IR1 and IR2.
The location of the partitioning site par and the binding sites
for the host protein DnaA are also shown.
In plasmid pSC101 and many of the broadhost-range plasmids, the ori region contains three
to seven copies of an iteron sequence which is 17 to
22 bp long. Close to the ori region there is a gene,
called repA in pSC101, which encodes the RepA
protein. This protein, which is the only plasmidencoded protein required for replication, binds to the
iterons and initiates DNA synthesis (Fig. 4.5).
Copy-number control is exerted by two superimposed mechanisms. First, the RepA protein represses
its own synthesis by binding to its own promoter
region and blocking transcription of its own gene
(Ingmer & Cohen 1993). If the copy number is high,
synthesis of RepA will be repressed. After cell division, the copy number and concentration of RepA
will drop and replication will be initiated. Mutations
in the RepA protein can lead to increased copy number (Ingmer & Cohen 1993, Cereghino et al. 1994).
Secondly, the RepA protein can link two plasmids
together, by binding to their iteron sequences, thereby preventing them from initiating replication. By
this mechanism, known as handcuffing (McEachern
et al. 1989), the replication of iteron plasmids will
depend both on the concentration of RepA protein
and on the concentration of the plasmids themselves.
Partitioning and segregative
stability of plasmids
The loss of plasmids due to defective partitioning
is called segregative instability. Naturally occurring
47
plasmids are stably maintained because they contain a partitioning function, par, which ensures that
they are stably maintained at each cell division.
Such par regions are essential for stability of lowcopy-number plasmids (for review, see Bingle &
Thomas 2001). The higher-copy-number plasmid
Col E1 also contains a par region but this is deleted in
many Col E1-derived cloning vectors, e.g. pBR322.
Although the copy number of vectors such as pBR322
is usually high, plasmid-free cells arise under nutrient limitation or other stress conditions ( Jones et al.
1980, Nugent et al. 1983). The par region from a
plasmid such as pSC101 can be cloned into pBR322,
thereby stabilizing the plasmid (Primrose et al. 1983).
DNA superhelicity is involved in the partitioning
mechanism (Miller et al. 1990). pSC101 derivatives
lacking the par locus show decreased overall superhelical density as compared with wild-type pSC101.
Partition-defective mutants of pSC101 and similar
mutants of unrelated plasmids are stabilized in
Eseherichia coli by topA mutations, which increase
negative DNA supercoiling. Conversely, DNA gyrase
inhibitors and mutations in DNA gyrase increase
the rate of loss of par-defective pSC101 derivatives.
Plasmid instability may also arise due to the formation of multimeric forms of a plasmid. The mechanism that controls the copy number of a plasmid
ensures a fixed number of plasmid origins per
bacterium. Cells containing multimeric plasmids
have the same number of plasmid origins but fewer
plasmid molecules, which leads to segregative instability if they lack a partitioning function. These multimeric forms are not seen with Col E1, which has
a natural method of resolving multimers back to
monomers. It contains a highly recombinogenic site
(cer). If the cer sequence occurs more than once in a
plasmid, as in a multimer, the host-cell Xer protein
promotes recombination, thereby regenerating monomers (Summers & Sherratt 1984, Guhathakurta
et al. 1996; for review, see Summers 1998).
Incompatibility of plasmids
Plasmid incompatibility is the inability of two different plasmids to coexist in the same cell in the
absence of selection pressure. The term incompatibility can only be used when it is certain that entry of
the second plasmid has taken place and that DNA
POGC04 9/11/2001 10:59 AM Page 48
48
CHAPTER 4
restriction is not involved. Groups of plasmids which
are mutually incompatible are considered to belong
to the same incompatibility (Inc) group. Over 30 incompatibility groups have been defined in E. coli and
13 for plasmids of S. aureus. Plasmids will be incompatible if they have the same mechanism of replication control. Not surprisingly, by changing the
sequence of the RNA I/RNA II region of plasmids
with antisense control of copy number, it is possible
to change their incompatibility group. Alternatively,
they will be incompatible if they share the same par
region (Austin & Nordstrom 1990, Firsheim & Kim
1997).
Upper band containing
chromosomal DNA and
open plasmid circles
The purification of plasmid DNA
An obvious prerequisite for cloning in plasmids is
the purification of the plasmid DNA. Although a
wide range of plasmid DNAs are now routinely
purified, the methods used are not without their
problems. Undoubtedly the trickiest stage is the lysis
of the host cells; both incomplete lysis and total
dissolution of the cells result in greatly reduced
recoveries of plasmid DNA. The ideal situation
occurs when each cell is just sufficiently broken to
permit the plasmid DNA to escape without too much
contaminating chromosomal DNA. Provided the
lysis is done gently, most of the chromosomal DNA
released will be of high molecular weight and can be
removed, along with cell debris, by high-speed
centrifugation to yield a cleared lysate. The production of satisfactory cleared lysates from bacteria
other than E. coli, particularly if large plasmids are to
be isolated, is frequently a combination of skill, luck
and patience.
Many methods are available for isolating pure
plasmid DNA from cleared lysates but only two will
be described here. The first of these is the ‘classical’
method and is due to Vinograd (Radloff et al. 1967).
This method involves isopycnic centrifugation of
cleared lysates in a solution of CsCl containing
ethidium bromide (EtBr). EtBr binds by intercalating
between the DNA base pairs, and in so doing causes
the DNA to unwind. A CCC DNA molecule, such as
a plasmid, has no free ends and can only unwind to
a limited extent, thus limiting the amount of EtBr
bound. A linear DNA molecule, such as fragmented
chromosomal DNA, has no such topological con-
Lower band of
covalently closed
circular plasmid DNA
Fig. 4.6 Purification of Col E1 KanR plasmid DNA by
isopycnic centrifugation in a CsCl–EtBr gradient.
(Photograph by courtesy of Dr G. Birnie.)
straints and can therefore bind more of the EtBr
molecules. Because the density of the DNA–EtBr
complex decreases as more EtBr is bound, and
because more EtBr can be bound to a linear molecule
than to a covalent circle, the covalent circle has a
higher density at saturating concentrations of EtBr.
Thus covalent circles (i.e. plasmids) can be separated
from linear chromosomal DNA (Fig. 4.6).
Currently the most popular method of extracting
and purifying plasmid DNA is that of Birnboim and
Doly (1979). This method makes use of the observation that there is a narrow range of pH (12.0–12.5)
within which denaturation of linear DNA, but not
covalently closed circular DNA, occurs. Plasmidcontaining cells are treated with lysozyme to weaken
the cell wall and then lysed with sodium hydroxide
and sodium dodecyl sulphate (SDS). Chromosomal
DNA remains in a high-molecular-weight form but
is denatured. Upon neutralization with acidic sodium
acetate, the chromosomal DNA renatures and aggregates to form an insoluble network. Simultaneously,
POGC04 9/11/2001 10:59 AM Page 49
Biology of plasmid and phage vectors
the high concentration of sodium acetate causes
precipitation of protein–SDS complexes and of highmolecular-weight RNA. Provided the pH of the
alkaline denaturation step has been carefully controlled, the CCC plasmid DNA molecules will remain
in a native state and in solution, while the contaminating macromolecules co-precipitate. The precipitate
can be removed by centrifugation and the plasmid
concentrated by ethanol precipitation. If necessary,
the plasmid DNA can be purified further by gel
filtration.
Recently a number of commercial suppliers of convenience molecular-biology products have developed
kits to improve the yield and purity of plasmid DNA.
All of them take advantage of the benefits of alkaline
lysis and have as their starting-point the cleared
lysate. The plasmid DNA is selectively bound to an
ion-exchange material, prepacked in columns or
tubes, in the presence of a chaotropic agent (e.g.
guanidinium hydrochloride). After washing away
the contaminants, the purified plasmid is eluted in a
small volume.
The yield of plasmid is affected by a number of
factors. The first of these is the actual copy number inside the cells at the time of harvest. The copynumber control systems described earlier are not the
only factors affecting yield. The copy number is also
affected by the growth medium, the stage of growth
and the genotype of the host cell (Nugent et al. 1983,
Seelke et al. 1987, Duttweiler & Gross 1998). The
second and most important factor is the care taken
in making the cleared lysate. Unfortunately, the
commercially available kits have not removed the
vagaries of this procedure. Finally, the presence in
the host cell of a wild-type endA gene can affect the
recovery of plasmid. The product of the endA gene
is endonuclease I, a periplasmic protein whose
substrate is double-stranded DNA. The function
of endonuclease I is not fully understood. Strains
bearing endA mutations have no obvious phenotype
other than improved stability and yield of plasmid
obtained from them.
Although most cloning vehicles are of low molecular weight (see next section), it is sometimes necessary to use the much larger conjugative plasmids.
Although these high-molecular-weight plasmids
can be isolated by the methods just described, the
yields are often very low. Either there is inefficient
49
release of the plasmids from the cells as a consequence of their size or there is physical destruction
caused by shear forces during the various manipulative steps. A number of alternative procedures have
been described (Gowland & Hardmann 1986), many
of which are a variation on that of Eckhardt (1978).
Bacteria are suspended in a mixture of Ficoll and
lysozyme and this results in a weakening of the cell
walls. The samples are then placed in the slots of an
agarose gel, where the cells are lysed by the addition
of detergent. The plasmids are subsequently extracted
from the gel following electrophoresis. The use of
agarose, which melts at low temperature, facilitates
extraction of the plasmid from the gel.
Desirable properties of
plasmid cloning vehicles
An ideal cloning vehicle would have the following
three properties:
• low molecular weight;
• ability to confer readily selectable phenotypic traits
on host cells;
• single sites for a large number of restriction
endonucleases, preferably in genes with a readily
scorable phenotype.
The advantages of a low molecular weight are
several. First, the plasmid is much easier to handle,
i.e. it is more resistant to damage by shearing, and
is readily isolated from host cells. Secondly, lowmolecular-weight plasmids are usually present as
multiple copies (see Table 4.2), and this not only
facilitates their isolation but leads to gene dosage
effects for all cloned genes. Finally, with a low
molecular weight there is less chance that the vector
will have multiple substrate sites for any restriction
endonuclease (see below).
After a piece of foreign DNA is inserted into a
vector, the resulting chimeric molecules have to
be transformed into a suitable recipient. Since the
efficiency of transformation is so low, it is essential
that the chimeras have some readily scorable
phenotype. Usually this results from some gene, e.g.
antibiotic resistance, carried on the vector, but could
also be produced by a gene carried on the inserted
DNA.
One of the first steps in cloning is to cut the vector
DNA and the DNA to be inserted with either the
POGC04 9/11/2001 10:59 AM Page 50
50
CHAPTER 4
same endonuclease or ones producing the same
ends. If the vector has more than one site for the
endonuclease, more than one fragment will be
produced. When the two samples of cleaved DNA
are subsequently mixed and ligated, the resulting
chimeras will, in all probability, lack one of the
vector fragments. It is advantageous if insertion of
foreign DNA at endonuclease-sensitive sites inactivates a gene whose phenotype is readily scorable,
for in this way it is possible to distinguish chimeras
from cleaved plasmid molecules which have selfannealed. Of course, readily detectable insertional
inactivation is not essential if the vector and insert
are to be joined by the homopolymer tailing method
(see p. 40) or if the insert confers a new phenotype
on host cells.
(a)
pBR322, a purpose-built cloning vehicle
In early cloning experiments, the cloning vectors
used were natural plasmids, such as Col E1 and
pSC101. While these plasmids are small and have
single sites for the common restriction endonucleases, they have limited genetic markers for selecting
transformants. For this reason, considerable effort
was expended on constructing, in vitro, superior
cloning vectors. The best, and most widely used of
these early purpose-built vectors is pBR322. Plasmid
pBR322 contains the ApR and TcR genes of RSF2124
and pSC101, respectively, combined with replication
elements of pMB1, a Col E1-like plasmid (Fig. 4.7a).
The origins of pBR322 and its progenitor, pBR313,
are shown in Fig. 4.7b, and details of its construction
can be found in the papers of Bolivar et al. (1977a,b).
(b)
RSF 2124-derived
material
1
R1 drd 19
ApR CmR SmR
SuR KmR
4361/1
pSC101-derived
material
EcoRI
pMB1
2
ColE1
R
A
R7268
ApR
p
5
R
Tc
pMB3
ApR
pSF2124
ApR
3145
Or
3
pSC101
TcR
igin
1763
pMB1-derived material
pMB8
4
6
pMB9
TcR
6
pBR312
ApR TcR
7
pBR313
ApR TcR
8
pBR322
ApR TcR
Fig. 4.7 The origins of plasmid pBR322. (a) The boundaries between the pSC101, pMB1 and RSF2124-derived material. The
numbers indicate the positions of the junctions in base pairs from the unique EcoRI site. (b) The molecular origins of plasmid
pBR322. R7268 was isolated in London in 1963 and later renamed R1. 1, A variant, R1drd19, which was derepressed for mating
transfer, was isolated. 2, The ApR transposon, Tn3, from this plasmid was transposed on to pMB1 to form pMB3. 3, This plasmid
was reduced in size by EcoRI* rearrangement to form a tiny plasmid, pMB8, which carries only colicin immunity. 4, EcoRI*
fragments from pSC101 were combined with pMB8 opened at its unique EcoRI site and the resulting chimeric molecule
rearranged by EcoRI* activity to generate pMB9. 5, In a separate event, the Tn3 of R1drd19 was transposed to Col E1 to form
pSF2124. 6, The Tn3 element was then transposed to pMB9 to form pBR312. 7, EcoRI* rearrangement of pBR312 led to the
formation of pBR313, from which (8) two separate fragments were isolated and ligated together to form pBR322. During this
series of constructions, R1 and Col E1 served only as carries for Tn3. (Reproduced by courtesy of Dr G. Sutcliffe and Cold Spring
Harbor Laboratory.)
POGC04 9/11/2001 10:59 AM Page 51
Biology of plasmid and phage vectors
Plasmid pBR322 has been completely sequenced.
The original published sequence (Sutcliffe 1979) was
4362 bp long. Position O of the sequence was arbitrarily set between the A and T residues of the EcoRI
recognition sequence (GAATTC). The sequence was
revised by the inclusion of an additional CG base
pair at position 526, thus increasing the size of the
plasmid to 4363 bp (Backman & Boyer 1983, Peden
1983). More recently, Watson (1988) has revised the
size yet again, this time to 4361 bp, by eliminating
base pairs at coordinates 1893 and 1915. The most
useful aspect of the DNA sequence is that it totally
characterizes pBR322 in terms of its restriction sites,
such that the exact length of every fragment can be
calculated. These fragments can serve as DNA markers
for sizing any other DNA fragment in the range of
several base pairs up to the entire length of the plasmid.
There are over 40 enzymes with unique cleavage
sites on the pBR322 genome (Fig. 4.8). The target
sites of 11 of these enzymes lie within the TcR gene,
and there are sites for a further two (ClaI and HindIII)
within the promoter of that gene. There are unique
sites for six enzymes within the ApR gene. Thus,
cloning in pBR322 with the aid of any one of those
19 enzymes will result in insertional inactivation of
either the ApR or the TcR markers. However, cloning
in the other unique sites does not permit the easy
selection of recombinants, because neither of the
antibiotic resistance determinants is inactivated.
Following manipulation in vitro, E. coli cells transformed with plasmids with inserts in the TcR gene
can be distinguished from those cells transformed
with recircularized vector. The former are ApR and
TcS, whereas the latter are both ApR and TcR. In
practice, transformants are selected on the basis
of their Ap resistance and then replica-plated on to
Tc-containing media to identify those that are TcS.
Cells transformed with pBR322 derivatives carrying
inserts in the ApR gene can be identified more readily
(Boyko & Ganschow 1982). Detection is based upon
the ability of the β-lactamase produced by ApR cells
to convert penicillin to penicilloic acid, which in turn
binds iodine. Transformants are selected on rich
medium containing soluble starch and Tc. When
colonized plates are flooded with an indicator solution of iodine and penicillin, β-lactamase-producing
(ApR) colonies clear the indicator solution whereas
ApS colonies do not.
51
The PstI site in the ApR gene is particularly useful,
because the 3′ tetranucleotide extensions formed on
digestion are ideal substrates for terminal transferase.
Thus this site is excellent for cloning by the homopolymer tailing method described in the previous
chapter (see p. 40). If oligo(dG.dC) tailing is used, the
PstI site is regenerated (see Fig. 3.11) and the insert
may be cut out with that enzyme.
Plasmid pBR322 has been a widely used cloning
vehicle. In addition, it has been widely used as a
model system for the study of prokaryotic transcription and translation, as well as investigation of the
effects of topological changes on DNA conformation.
The popularity of pBR322 is a direct result of the
availability of an extensive body of information on
its structure and function. This in turn is increased
with each new study. The reader wishing more
detail on the structural features, transcriptional signals, replication, amplification, stability and conjugal mobility of pBR322 should consult the review
of Balbás et al. (1986).
Example of the use of plasmid pBR322 as a vector:
isolation of DNA fragments which carry promoters
Cloning into the HindIII site of pBR322 generally
results in loss of tetracycline resistance. However,
in some recombinants, TcR is retained or even
increased. This is because the HindIII site lies within
the promoter rather than the coding sequence.
Thus whether or not insertional inactivation occurs depends on whether the cloned DNA carries a
promoter-like sequence able to initiate transcription
of the TcR gene. Widera et al. (1978) have used
this technique to search for promoter-containing
fragments.
Four structural domains can be recognized within
E. coli promoters. These are:
• position 1, the purine initiation nucleotide from
which RNA synthesis begins;
• position −6 to −12, the Pribnow box;
• the region around base pair −35;
• the sequence between base pairs −12 and −35.
Although the HindIII site lies within the Pribnow
box (Rodriguez et al. 1979) the box is re-created on
insertion of a foreign DNA fragment. Thus when
insertional inactivation occurs it must be the region
from −13 to −40 which is modified.
POGC04 9/11/2001 05:13 PM Page 52
52
CHAPTER 4
Cla I - BspD I 23
Hind III 29
EcoR V 185
EcoRI - Apo I 4359
Aat II 4284
Nhe I 229
BsrB - Bci V I 4209
BamH I 375
Ssp I 4168
Ear I 4155
SgrA I 409
Ban II 471
Ban II 485
Dsa I 528
Eco57 I 4054
Xmn I 3961
Hinc II 3905
Sph I 562
Sca I 3844
EcoN I 622
Sal I 651
Pvu I 3733
Hinc II 651
Pst I 3607
Acc I 651
Ap
Tc
BsrD I 3608
PshA I 712
Ase I 3537
Eag I 939
Bsa I 3433
BsrD I 3420
Nru I 972
Ahd I 3361
BstAP I 1045
†
BspM I 1063
PflM I 1315
Bsm I 1353
Sty I 1369
Eco57 I 3000
I
OR
PflM I 1364
AlwN I 2884
Ava I - BsoB I 1425
PpuM I 1438
Msc I 1444
Bci V I 2682
Drd I 2575
Dsa I 1447
ROP
Afl III - BspLU11 I 2473
BsrB I 2408
Ear I 2351
Sap I 2350
Nde I 2295
BstAP I 2291
BstZ17 I - Acc I 2244
Bsa A I 2225
PpuM I 1480
Bsg I 1656
Xmn I 2029
Pvu II 2064
BspE I 1664
BsaB I 1668
BsmB I 2122
Drd I 2162
PflFI - Tth111 I 2217
Fig. 4.8 Restriction map of plasmid pBR322 showing the location and direction of transcription of the ampicillin (Ap) and
tetracycline (Tc) resistance loci, the origin of replication (ori) and the Col E1-derived Rop gene. The map shows the restriction
sites of those enzymes that cut the molecule once or twice. The unique sites are shown in bold type. The coordinates refer to the
position of the 5′ base in each recognition sequence with the first T in the EcoRI site being designated as nucleotide number 1. The
exact positions of the loci are: Tc, 86–1268; Ap, 4084–3296; Rop, 1918–2105 and the origin of replication, 2535.
POGC04 9/11/2001 10:59 AM Page 53
Biology of plasmid and phage vectors
Improved vectors derived from pBR322
Over the years, numerous different derivatives of
pBR322 have been constructed, many to fulfil
special-purpose cloning needs. A compilation of the
properties of some of these plasmids has been provided by Balbás et al. (1986).
Much of the early work on the improvement of
pBR322 centred on the insertion of additional unique
restriction sites and selectable markers, e.g. pBR325
encodes chloramphenicol resistance in addition
to ampicillin and tetracycline resistance and has a
unique EcoRI site in the CmR gene. Initially, each
new vector was constructed in a series of steps
analogous to those used in the generation of
pBR322 itself (Fig. 4.7). Then the construction of
improved vectors was simplified (Vieira & Messing
1982, 1987, Yanisch-Perron et al. 1985) by the use
of polylinkers or multiple cloning sites (MCS), as
exemplified by the pUC vectors (Fig. 4.9). An MCS is
a short DNA sequence, 2.8 kb in the case of pUC19,
carrying sites for many different restriction endonucleases. An MCS increases the number of potential
cloning strategies available by extending the range
of enzymes that can be used to generate a restriction
fragment suitable for cloning. By combining them
within an MCS, the sites are made contiguous, so
that any two sites within it can be cleaved simultaneously without excising vector sequences.
The pUC vectors also incorporate a DNA sequence
that permits rapid visual detection of an insert.
The MCS is inserted into the lacZ′ sequence, which
encodes the promoter and the α-peptide of βgalactosidase. The insertion of the MCS into the lacZ′
fragment does not affect the ability of the α-peptide
to mediate complementation, but cloning DNA fragments into the MCS does. Therefore, recombinants
can be detected by blue/white screening on growth
medium containing Xgal (see Box 3.2 on p. 35). The
usual site for insertion of the MCS is between
the iniator ATG codon and codon 7, a region that
encodes a functionally non-essential part of the
α-complementation peptide. Recently, Slilaty and
Lebel (1998) have reported that blue/white colour
selection can be variable. They have found that
reliable inactivation of complementation occurs
only when the insert is made between codons 11
and 36.
53
Bacteriophage l
Essential features
Bacteriophage λ is a genetically complex but very
extensively studied virus of E. coli (Box 4.1). Because
it has been the object of so much moleculargenetical research, it was natural that, right from
the beginnings of gene manipulation, it should have
been investigated and developed as a vector. The
DNA of phage λ, in the form in which it is isolated
from the phage particle, is a linear duplex molecule
of about 48.5 kbp. The entire DNA sequence has
been determined (Sanger et al. 1982). At each
end are short single-stranded 5′ projections of 12
nucleotides, which are complementary in sequence
and by which the DNA adopts a circular structure
when it is injected into its host cell, i.e. λ DNA
naturally has cohesive termini, which associate
to form the cos site.
Functionally related genes of phage λ are clustered together on the map, except for the two positive regulatory genes N and Q. Genes on the left of the
conventional linear map (Fig. 4.10) code for head
and tail proteins of the phage particle. Genes of the
central region are concerned with recombination
(e.g. red ) and the process of lysogenization, in which
the circularized chromosome is inserted into its host
chromosome and stably replicated along with it as
a prophage. Much of this central region, including
these genes, is not essential for phage growth and
can be deleted or replaced without seriously impairing the infectious growth cycle. Its dispensability
is crucially important, as will become apparent
later, in the construction of vector derivatives of
the phage. To the right of the central region are
genes concerned with regulation and prophage
immunity to superinfection (N, cro, cI), followed by
DNA synthesis (O, P), late function regulation (Q)
and host cell lysis (S, R). Figure 4.11 illustrates the
λ life cycle.
Promoters and control circuits
As we shall see, it is possible to insert foreign DNA
into the chromosome of phage-λ derivative and, in
some cases, foreign genes can be expressed efficiently via λ promoters. We must therefore briefly
POGC04 9/11/2001 10:59 AM Page 54
1
2
3
4
1
2
3
4
5
6
7
8
9 10 11 12 13 14 15 16 17 18 5
6
7
8
THR MET ILE THR pro ser leu his ala cys arg ser thr leu glu asp pro arg val pro ser ser ASN SER LEU ALA
ATG ACC ATG ATT ACG CGA AGC TTG CAT GCC TGC AGG TCG ACT CTA GAG GAT CCC CGG GTA CCG AGC TCG AAT TCA CTG GCC
HindIII
SphI
PsfI
SalI
XbaI
BamHI
KpnI
SstI
EcoRI
HaeIII
AccI
XmaI
HincII
SmaI
1
2
3
4
5
6
1
2
3
4
5
6
7
8
9 10 11 12 13 14 15 16 17 18 7
8
THR MET ILE THR ASN SER ser ser val pro gly asp pro leu glu ser thr cys arg his ala ser leu ala LEU ALA
pUC18
ATG ACC ATG ATT ACG AAT TCG AGC TCG GTA CCC GGG GAT CCT CTA GAG TCG ACC TGC AGG CAT GCA AGC TTG GCA CTG GCC
EcoRI
SstI
KpnI
BamHI
XbaI
SalI
PstI
SphI
HindIII
HaeIII
AccI
XmaI
HincII
SmaI
1
2
3
4
1
2
3
4
5
6
7
8
9 10 11 12 13 14 15 16 5
6
7
8
THR MET ILE THR pro ser leu gly cys arg ser thr leu glu asp pro arg ala ser ser ASN SER LEU ALA
pUC13
ATG ACC ATG ATT ACG CCA AGC TTG GGC TGC AGG TCG ACT CTA GAG GAT CCC CGG GCG AGC TCG AAT TCA CTG GCC
HindIII
PstI
SalI
XbaI
BamHI
SstI
EcoRI
HaeIII
AccI
XmaI
HincII
SmaI
3
1
2
3
4
5
6
1
2
4
5
6
7
8
9 10 11 12 13 14 15 16 17 18
THR MET ILE THR ASN SER ser ser pro gly asp pro leu glu ser thr cys ser pro ser leu ala LEU ALA
pUC12
ATG ACC ATG ATT ACG AAT TCG AGC TCG CCC GGG GAT CCT CTA GAG TCG ACC TGC AGC CCA AGC TTG GCA CTG GCC
SstI
BamHI
XbaI
SalI
PstI
HindIII
HaeIII
EcoRI
AccI
XmaI
HincII
SmaI
pUC19
1
2
3
4
1
2
3
4
5
6
7
8
9 10 11 5
6
7
8
THR MET ILE THR pro ser leu ala ala gly arg arg ile pro gly ASN SER LEU ALA
ATG ACC ATG ATT ACG CCA AGC TTG GCT GCA GGT CGA CGG ATC CCC GGG AAT TCA CTG GCC
EcoRI
HaeIII
HindIII
PstI
SalI
BamHI
AccI
XmaI
HincII
SmaI
1
2
3
4
5
6
1
2
3
4
5
6
7
8
9 10 11 7
8
THR MET ILE THR ASN SER arg gly ser val asp leu gln pro ser leu ala LEU ALA
pUC8
ATG ACC ATG ATT ACG AAT TCC CGG GGA TCC GTC GAC CTG CAG CCA AGC TTG GCA CTG GCC
EcoRI
BamHI
SalI
PstI
HindIII
HaeIII
AccI
XmaI
HincII
SmaI
1
2
3
4
5
1
2
3
4
5
6
7
8
9 10 11 12 13 14 6
THR MET ILE THR ASN ser pro asp pro ser thr cys arg ser thr asp pro gly asn SER
pUC7
ATG ACC ATG ATT ACG AAT TCC CCG GAT CCG TCG ACC TGC AGG TCG ACG GAT CCG GGG AAT TCA
PstI
BamHI
EcoRI
SalI
EcoRI
BamHI
SalI
AccI
AccI
HincII
HincII
pUC9
BsmB I 51 Nde I 183
BstAP I 179
Drd I 91
Nar I- Kas I-Sfo I 235
Bgl I 245
Fsp I 256
Pvu I 276
lac
Pvu II 306
Z
BsmB I 2683
EcoO 109 I 2674
Aat II 2617
Bci V I 2542
Ssp I 2501
Eco57 I 2381
Acl I 2297
Xmn I 2294
Polylinker
cloning sites
396-454
Bcg I 2215
Sca I 2177
Pvu II 628
Tfi I 641
Sap I 683
AP
Pvu I 2066
Ava II 2059
BsrD I 1935
Acl I 1924
Fsp I 1919
Tfi I 781
Afl III 806
Ava II 1837
Bgl I 1813
Bpm I 1784
BsrF I 1779
Bsa I 1766
BsrD I 1753
Drd I 908
Bci V I 1015
I
OR
Ahd I 1694
Eco57 I 1333
AlwN I 1217
Fig. 4.9 Genetic maps of some pUC plasmids. The multiple cloning site (MCS) is inserted into the lacZ gene but does not interfere
with gene function. The additional codons present in the lacZ gene as a result of the polylinker are labelled with lower-case letters.
These polylinker regions (MCS) are identical to those of the M13 mp series of vectors (see p. 63).
POGC04 9/11/2001 10:59 AM Page 55
Biology of plasmid and phage vectors
55
Box 4.1 Bacteriophage l: its important place in molecular biology and
recombinant DNA technology
DNA
Head
proteins
Tail proteins
In the early 1950s, following some initial studies on
Bacillus megaterium, André Lwoff and his colleagues
at the Institut Pasteur described the phenomenon of
lysogeny in E. coli. It became clear that certain strains
of E. coli were lysogenized by phage, that is to say,
these bacteria harboured phage l in a dormant
form, called a prophage. The lysogenic bacteria grew
normally and might easily not have been recognized
as lysogenic. However, when Lwoff exposed the
bacteria to a moderate dose of ultraviolet light, the
bacteria stopped growing, and after about 90 min
of incubation the bacteria lysed, releasing a crop
of phage into the medium.
The released phage were incapable of infecting
more E. coli than had been lysogenized by phage
l (this is called immunity to superinfection), but
non-lysogenic bacteria could be infected to yield
consider the promoters and control circuits affecting
λ gene expression (see Ptashne (1992) for an excellent monograph on phage-λ control circuits).
In the lytic cycle, λ transcription occurs in three
temporal stages: early, middle and late. Basically,
early gene transcription establishes the lytic cycle
another crop of virus. Not every non-lysogenic
bacterium yielded virus; some bacteria were
converted into lysogens because the phage
switched to the dormant lifestyle – becoming
prophage – rather than causing a lytic infection.
By the mid-1950s it was realized that the
prophage consisted of a phage l genome that had
become integrated into the E. coli chromosome. It
was also apparent to Lwoff’s colleagues, Jacob and
Monod, that the switching between the two states of
the virus – the lytic and lysogenic lifestyles – was an
example of a fundamental aspect of genetics that
was gaining increasing attention, gene regulation.
Intensive genetic and molecular biological analysis
of the phage, mainly in the 1960s and 1970s, led to
a good understanding of the virus. The key molecule
in maintaining the dormancy of the prophage and in
conferring immunity to superinfection is the phage
repressor, which is the product of the phage cI gene.
In 1967 the phage repressor was isolated by Mark
Ptashne (Ptashne 1967a,b). The advanced molecular
genetics of the phage made it a good candidate for
development as a vector, beginning in the 1970s and
continuing to the present day, as described in the
text. The development of vectors exploited the fact
that a considerable portion of the phage genome
encodes functions that are not needed for the
infectious cycle. The ability to package recombinant
phage DNA into virus particles in vitro was an
important development for library construction
(Hohn & Murray 1977).
A landmark in molecular biology was reached
when the entire sequence of the phage l genome,
48 502 nucleotide pairs, was determined by Fred
Sanger and his colleagues (Sanger et al. 1982).
(in competition with lysogeny), middle gene products replicate and recombine the DNA and late
gene products package this DNA into mature phage
particles. Following infection of a sensitive host,
early transcription proceeds from major promoters
situated immediately to the left (PL) and right (PR) of
POGC04 9/11/2001 10:59 AM Page 56
CHAPTER 4
b2
reg
ion
Int
rec egrat
om ion
bin an
atio d
Reg
n
im ulati
mu on
nit
a
nd
y
DN
As
Lat
e fu ynth
esi
nct
s
Ho ion r
e
st l
ysi gula
tio
s
n
56
Head
Left
cohesive
end
%λ 1
Tail
AWB DEFZ
10
20
xis
cro
att int red N cI
OP Q SR
gam
J
30
40
50
60
70
80
90
Right
cohesive
end
100 %λ
Non-essential region
Fig. 4.10 Map of the λ chromosome,
showing the physical position of some
genes on the full-length DNA of wildtype bacteriophage λ. Clusters of
functionally related genes are indicated.
Absorption
R
A
A
Cell envelope
Host lysis
R
Addition of
phage tails
A
Cleavage
Nicks sealed
by ligase
cos
cos
cos
R
Early
replication
(bidirectional
mode)
A
Lysogeny
cos
R
A
Origin
cos
R
cos
gal
Recombination
att
x
att
E.coli chromosome
cos
Integration
Late replication
(rolling circle mode)
bio
gal N
R cos A
att
Excision
J
bio
att
Replication along with
E.coli chromosome
the repressor gene (cI ) (Fig. 4.12). This transcription
is subject to repression by the product of the cI gene
and in a lysogen this repression is the basis of
immunity to superinfecting λ. Early in infection,
transcripts from PL and PR stop at termination sites tL
and tR1. The site tR2 stops any transcripts that escape
beyond tR1. Lambda switches from early- to middle-
Fig. 4.11 Replication of phage-λ DNA in
lytic and lysogenic cycles.
stage transcription by anti-termination. The N gene
product, expressed from PL, directs this switch. It
interacts with RNA polymerase and, antagonizing
the action of host termination protein ρ, permits it to
ignore the stop signals so that PL and PR transcripts
extend into genes such as red, O and P necessary for
the middle stage. The early and middle transcripts
POGC04 9/11/2001 10:59 AM Page 57
Biology of plasmid and phage vectors
PL PR
tL
N
tR1
cl
O
red
tR2
P
int
Q
P′R
S
R
cos
J
A
Z F
Legend
Early transcripts
Middle transcripts
Late transcripts
Fig. 4.12 Major promoters and transcriptional termination
sites of phage λ. (See text for details.)
and patterns of expression therefore overlap. The cro
product, when sufficient has accumulated, prevents
transcription from PL and PR. The gene Q is expressed from the distal portion of the extended PR
transcript and is responsible for the middle-to-late
switch. This also operates by anti-termination. The
Q product specifically anti-terminates the short PR
transcript, extending it into the late genes, across
the cohered cos region, so that many mature phage
particles are ultimately produced.
Both N and Q play positive regulatory roles essential for phage growth and plaque formation; but an
N − phage can produce a small plaque if the termination site tR2 is removed by a small deletion termed nin
(N-independent) as in λN− nin.
Vector DNA
Wild-type λ DNA contains several target sites for
most of the commonly used restriction endonucleases and so is not itself suitable as a vector.
Derivatives of the wild-type phage have therefore
been produced that either have a single target site
at which foreign DNA can be inserted (insertional
vectors) or have a pair of sites defining a fragment
57
that can be removed (stuffer) and replaced by foreign
DNA (replacement vectors). Since phage λ can accommodate only about 5% more than its normal complement of DNA, vector derivatives are constructed
with deletions to increase the space within the
genome. The shortest λ DNA molecules that produce
plaques of nearly normal size are 25% deleted.
Apparently, if too much non-essential DNA is
deleted from the genome, it cannot be packaged into
phage particles efficiently. This can be turned to
advantage for, if the replaceable fragment of a
replacement-type vector is either removed by physical separation or effectively destroyed by treatment
with a second restriction endonuclease that cuts it
alone, then the deleted vector genome can give rise
to plaques only if a new DNA segment is inserted into
it. This amounts to positive selection for recombinant phage carrying foreign DNA.
Many vector derivatives of both the insertional and
replacement types were produced by several groups
of researchers early in the development of recombinant DNA technology (e.g. Thomas et al. 1974, Murray
& Murray 1975, Blattner et al. 1977, Leder et al.
1977). Most of these vectors were constructed for
use with EcoRI, BamHI or HindIII, but their application could be extended to other endonucleases by
the use of linker molecules. These early vectors have
been largely superseded by improved vectors for
rapid and efficient genomic or complementary DNA
(cDNA) library construction (see Chapter 6).
λ vectors
Improved phage-λ
As with plasmid vectors, improved phage-vector
derivatives have been developed. There have been
several aims, among which are the following.
• To increase the capacity for foreign DNA fragments, preferably for fragments generated by any
one of several restriction enzymes (reviewed by
Murray 1983).
• To devise methods for positively selecting recombinant formation.
• To allow RNA probes to be conveniently prepared
by transcription of the foreign DNA insert; this
facilitates the screening of libraries in chromosome
walking procedures. An example of a vector with
this property is λZAP (see p. 93).
• To develop vectors for the insertion of eukaryotic
cDNA (p. 93) such that expression of the cDNA, in
POGC04 9/11/2001 10:59 AM Page 58
left arm
(20 kb)
EcoRI
BamHI
SalI
central stuffer
(14 kb)
right arm
(9 kb)
EMBL3 vector
left arm
(20 kb)
SalI
BamHI
EcoRI
the form of a fusion polypeptide with β-galactosidase,
is driven in E. coli; this form of expression vector is
useful in antibody screening. An example of such a
vector is λgt11.
The first two points will be discussed here. The discussion of improved vectors in library construction
and screening is deferred until Chapter 6.
The maximum capacity of phage-λ derivatives
can only be attained with vectors of the replacement
type, so that there has also been an accompanying
incentive to devise methods for positively selecting
recombinant formation without the need for prior
removal of the stuffer fragment. Even when steps are
taken to remove the stuffer fragment by physical purification of vector arms, small contaminating amounts
may remain, so that genetic selection for recombinant formation remains desirable. The usual method
of achieving this is to exploit the Spi− phenotype.
Wild-type λ cannot grow on E. coli strains lysogenic for phage P2; in other words, the λ phage is
Spi+ (sensitive to P2 inhibition). It has been shown
that the products of λ genes red and gam, which lie in
the region 64–69% on the physical map, are
responsible for the inhibition of growth in a P2 lysogen (Herskowitz 1974, Sprague et al. 1978, Murray
1983). Hence vectors have been derived in which
the stuffer fragment includes the region 64–69%, so
that recombinants in which this has been replaced
by foreign DNA are phenotypically Spi− and can be
positively selected by plating on a P2 lysogen (Karn
et al. 1986, Loenen & Brammar 1980).
Deletion of the gam gene has other consequences.
The gam product is necessary for the normal switch
in λ DNA replication from the bidirectional mode to
the rolling-circle mode (see Fig. 4.11). Gam− phage
cannot generate the concatemeric linear DNA
which is normally the substrate for packaging into
phage heads. However, gam− phage do form plaques
because the rec and red recombination systems act
on circular DNA molecules to form multimers, which
can be packaged. gam− red − phage are totally dependent upon rec-mediated exchange for plaque formation on rec+ bacteria. λ DNA is a poor substrate for
this rec-mediated exchange. Therefore, such phage
make vanishingly small plaques unless they contain
one or more short DNA sequences called chi (crossover hot-spot instigator) sites, which stimulate
rec-mediated exchange. Many of the current replacement vectors generate red − gam − clones and
SalI
BamHI
EcoRI
CHAPTER 4
EcoRI
BamHI
SalI
58
central stuffer
(14 kb)
right arm
(9 kb)
EMBL4 vector
5′. . . GGATC TGGGT CGACG GATCC GGGGA ATTCC CAGAT CC . . .3′
SalI
BamHI
EcoRI
Fig. 4.13 The structure of bacteriophage λ cloning vectors
EMBL3 and EMBL4. The polylinker sequence is present in
opposite orientation in the two vectors.
so have been constructed with a chi site within the
non-replaceable part of the phage.
The most recent generation of λ vectors, which
are based on EMBL3 and EMBL4 (Frischauf et al.
1983; Fig. 4.13), have a capacity for DNA of size
9–23 kb. As well as being chi+, they have polylinkers
flanking the stuffer fragment to facilitate library
construction. Phages with inserts can be selected on
the basis of their Spi− phenotype, but there is an
alternative. The vector can be digested with BamHI
and EcoRI prior to ligation with foreign DNA fragments produced with BamHI. If the small BamHI–
EcoRI fragments from the polylinkers are removed,
the stuffer fragment will not be reincorporated.
λ DNA in vitro
Packaging phage-λ
So far, we have considered only one way of introducing manipulated phage DNA into the host
bacterium, i.e. by transfection of competent bacteria
(see Chapter 2). Using freshly prepared λ DNA that
has not been subjected to any gene-manipulation
procedures, transfection will result typically in about
105 plaques/µg of DNA. In a gene-manipulation
experiment in which the vector DNA is restricted
and then ligated with foreign DNA, this figure
is reduced to about 104–103 plaques/µg of vector
DNA. Even with perfectly efficient nucleic acid
POGC04 9/11/2001 10:59 AM Page 59
Biology of plasmid and phage vectors
59
Head precursor
Major capsid protein
is product of gene E
Concatemeric DNA
cos
cos
cos
Endonucleolytic cleavage by product of
gene A at cos sites. At each cos site nicks
are introduced 12 bp apart on opposite
strands of the DNA, hence generating
the cohesive termini of λDNA as it is
found in the phage particle. ATP is
required for this process
Product of gene D
included in capsid
Assembly proteins, products of genes W, FII plus
completed tails
Mature particle
Fig. 4.14 Simplified scheme showing
packaging of phage-λ DNA into phage
particles.
biochemistry, some of this reduction is inevitable.
It is a consequence of the random association of
fragments in the ligation reaction, which produces
molecules with a variety of fragment combinations,
many of which are inviable. Yet, in some contexts,
106 or more recombinants are required. The scale of
such experiments can be kept within a reasonable
limit by packaging the recombinant DNA into
mature phage particles in vitro.
Placing the recombinant DNA in a phage coat
allows it to be introduced into the host bacteria by
the normal processes of phage infection, i.e. phage
adsorption followed by DNA injection. Depending
upon the details of the experimental design, packaging in vitro yields about 106 plaques/µg of vector
DNA after the ligation reaction.
Figure 4.14 shows some of the events occurring
during the packaging process that take place within
the host during normal phage growth and which we
now require to perform in vitro. Phage DNA in concatemeric form, produced by a rolling-circle replication mechanism (see Fig. 4.11), is the substrate for
the packaging reaction. In the presence of phage
head precursor (the product of gene E is the major
capsid protein) and the product of gene A, the
concatemeric DNA is cleaved into monomers and
encapsidated. Nicks are introduced in opposite
strands of the DNA, 12 nucleotide pairs apart at
each cos site, to produce the linear monomer with its
cohesive termini. The product of gene D is then
incorporated into what now becomes a completed
phage head. The products of genes W and FII,
among others, then unite the head with a separately
assembled tail structure to form the mature particle.
The principle of packaging in vitro is to supply the
ligated recombinant DNA with high concentrations
of phage-head precursor, packaging proteins and
phage tails. Practically, this is most efficiently performed in a very concentrated mixed lysate of
two induced lysogens, one of which is blocked at
the pre-head stage by an amber mutation in gene D
and therefore accumulates this precursor, while the
other is prevented from forming any head structure
by an amber mutation in gene E (Hohn & Murray
1977). In the mixed lysate, genetic complementation
occurs and exogenous DNA is packaged (Fig. 4.15).
Although concatemeric DNA is the substrate for
packaging (covalently joined concatemers are, of
course, produced in the ligation reaction by association of the natural cohesive ends of λ), the in vitro
system will package added monomeric DNA, which
presumably first concatemerizes non-covalently.
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60
CHAPTER 4
Phage genes
Phage genes
Assembly
proteins
Assembly
proteins
Induced λ lysogen E.coli
BHB2688 E amber mutation.
Produces λ tails, protein D,
assembly proteins, but no
protein E and hence no
pre-heads
Induced λ lysogen E.coli
BHB2690 D amber mutation.
Produces λ tails, pre-heads
including protein E, assembly
proteins, but DNA packaging
blocked by lack of protein D
pD
Lyse, mix
Add ATP and
concatemerized λDNA
COS
COS
+
COS
+
Assembly
proteins
COS
+
pD
Mature phage
particles
There are two potential problems associated with
packaging in vitro. First, endogenous DNA derived
from the induced prophages of the lysogens used to
prepare the packaging lysate can itself be packaged.
This can be overcome by choosing the appropriate
genotype for these prophages, i.e. excision upon
induction is inhibited by the b2 deletion (Gottesmann & Yarmolinsky 1968) and imm 434 immunity will prevent plaque formation if an imm 434
lysogenic bacterium is used for plating the complex
reaction mixture. Additionally, if the vector does not
contain any amber mutation a non-suppressing host
bacterium can be used so that endogenous DNA
will not give rise to plaques. The second potential
problem arises from recombination in the lysate
between exogenous DNA and induced prophage
markers. If troublesome, this can be overcome by
using recombination-deficient (i.e. red − rec −) lysogens and by UV-irradiating the cells used to prepare
the lysate, so eliminating the biological activity of
the endogenous DNA (Hohn & Murray 1977).
Fig. 4.15 In vitro packaging of concatemerized
phage-λ DNA in a mixed lysate.
DNA cloning with single-stranded
DNA vectors
M13, f1 and fd are filamentous coliphages containing a circular single-stranded DNA molecule. These
coliphages have been developed as cloning vectors,
for they have a number of advantages over other
vectors, including the other two classes of vector for
E. coli, plasmids and phage λ. However, in order to
appreciate their advantages, it is essential to have a
basic understanding of the biology of filamentous
phages.
The biology of the filamentous coliphages
The phage particles have dimensions 900 nm ×
9 nm and contain a single-stranded circular DNA
molecule, which is 6407 (M13) or 6408 (fd) nucleotides long. The complete nucleotide sequences of fd
and M13 are available and they are 97% homologous.
The differences consist mainly of isolated nucleotides
POGC04 9/11/2001 10:59 AM Page 61
Biology of plasmid and phage vectors
here and there, mostly affecting the redundant bases
of codons, with no blocks of sequence divergence.
Sequencing of f1 DNA indicates that it is very similar
to M13 DNA.
The filamentous phages only infect strains of
enteric bacteria harbouring F pili. The adsorption
site appears to be the end of the F pilus, but exactly
how the phage genome gets from the end of the F
pilus to the inside of the cell is not known.
Replication of phage DNA does not result in host-cell
lysis. Rather, infected cells continue to grow and
divide, albeit at a slower rate than uninfected cells,
and extrude virus particles. Up to 1000 phage particles may be released into the medium per cell per
generation (Fig. 4.16).
The single-stranded phage DNA enters the cell by
a process in which decapsidation and replication are
tightly coupled. The capsid proteins enter the cytoplasmic membrane as the viral DNA passes into the
cell while being converted to a double-stranded
replicative form (RF). The RF multiplies rapidly until
about 100 RF molecules are formed inside the cell.
Replication of the RF then becomes asymmetric, due
to the accumulation of a viral-encoded singlestranded specific DNA-binding protein. This protein
binds to the viral strand and prevents synthesis of
the complementary strand. From this point on, only
viral single strands are synthesized. These progeny
single strands are released from the cell as filamentous particles following morphogenesis at the cell
membrane. As the DNA passes through the membrane, the DNA-binding protein is stripped off and
replaced with capsid protein.
Why use single-stranded vectors?
For several applications of cloned DNA, singlestranded DNA is required. Sequencing by the original dideoxy method required single-stranded DNA,
as do techniques for oligonucleotide-directed mutagenesis and certain methods of probe preparation.
The use of vectors that occur in single-stranded form
is an attractive means of combining the cloning,
amplification and strand separation of an originally
double-stranded DNA fragment.
As single-stranded vectors, the filamentous phages
have a number of advantages. First, the phage DNA
is replicated via a double-stranded circular DNA
(RF) intermediate. This RF can be purified and
61
manipulated in vitro just like a plasmid. Secondly,
both RF and single-stranded DNA will transfect
competent E. coli cells to yield either plaques or
infected colonies, depending on the assay method.
Thirdly, the size of the phage particle is governed by
the size of the viral DNA and therefore there are no
packaging constraints. Indeed, viral DNA up to six
times the length of M13 DNA has been packaged
(Messing et al. 1981). Finally, with these phages it is
very easy to determine the orientation of an insert.
Although the relative orientation can be determined
from restriction analysis of RF, there is an easier
method (Barnes 1980). If two clones carry the insert
in opposite directions, the single-stranded DNA from
them will hybridize and this can be detected by
agarose gel electrophoresis. Phage from as little as
0.1 ml of culture can be used in assays of this sort,
making mass screening of cultures very easy.
In summary, as vectors, filamentous phages possess all the advantages of plasmids while producing
particles containing single-stranded DNA in an easily obtainable form.
Development of filamentous
phage vectors
Unlike λ, the filamentous coliphages do not have
any non-essential genes which can be used as
cloning sites. However, in M13 there is a 507 bp
intergenic region, from position 5498 to 6005 of the
DNA sequence, which contains the origins of DNA
replication for both the viral and the complementary
strands. In most of the vectors developed so far, foreign DNA has been inserted at this site, although it is
possible to clone at the carboxy-terminal end of gene
IV (Boeke et al. 1979). The wild-type phages are not
very promising as vectors because they contain very
few unique sites within the intergenic region: AsuI in
the case of fd, and AsuI and AvaI in the case of M13.
The first example of M13 cloning made use of one
of 10 BsuI sites in the genome, two of which are in
the intergenic region (Messing et al. 1977). For
cloning, M13 RF was partially digested with BsuI
and linear full-length molecules isolated by agarose
gel electrophoresis. These linear monomers were
blunt-end-ligated to a HindII restriction fragment
comprising the E. coli lac regulatory region and the
genetic information for the α-peptide of β-galactosidase. The complete ligation mixture was used to
POGC04 9/11/2001 10:59 AM Page 62
62
CHAPTER 4
Gene 8 protein
F pilus of F+ E.coli
Gene 3 protein
M13 filamentous phage particle
(870 × 6 nm). Single-stranded DNA
circle of 6407 nucleotides
(+)
5
2
RNA pol
4
3
DNA polIII
Nicking by gene
2 protein
5
6
Stage 1
Absorption and
conversion of
single-stranded
DNA to doublestranded
(replicative form,
RF) DNA
0–1 minute
post-infection
Stage 2
RF → RF rolling
circle replication
∼20 minutes
post-infection
8
(+)
7
Cutting by
gene 2 protein
Gene 5 protein
(+)
Stage 3
Gene 5 protein
switches DNA
synthesis to
production of
single strands
∼20 minutes
post-infection
to indefinitely
Virus particles extruded from
infected cell without lysis
Fig. 4.16 Life cycle and DNA replication of phage M13.
transform a strain of E. coli with a deletion of the βgalactosidase α-fragment and recombinant phage
detected by intragenic complementation on media
containing IPTG and Xgal (see Box 3.2 on p. 35).
One of the blue plaques was selected and the virus in
it designated M13 mp1.
Insertion of DNA fragments into the lac region of
M13 mp1 destroys its ability to form blue plaques,
POGC04 9/11/2001 10:59 AM Page 63
Biology of plasmid and phage vectors
Xmn I 357
X
BsrG I 1020
BsaB I 1148
SnaB I 1267
BspH I 1298
i
Cla I–BspD I 6881
Swa I 6782
Acl I 6772
Bpm I 6904
Dsa I 6616
BsmF I 6528
Bgl II 6934
Bsu36 I 6507
Bpm I 6492
Msl I 6996
Bgl I 6431
Fsp I 6425
Polylinker
Pvu I 6405
cloning sites
II
Ear I 6390
6229–6287
BsaH I 6000
cZ
a
L
Sfo I–Nar I–Kas I 6000
BsmB I 5970
Ava II 5914
BsoB I–Ava I 5824
Drd I 5758
Dra III 5715
Plus
Ban II 5642
ORI
Nae I–NgoM IV 5612
Minus
BsrF I 5612
63
La
c
I
VI
V
VII
IX
I
IV
Bsm I 1745
BsmF I 5089
III
BseR I 2007
Msc I 5079
AlwN I 2186
VI
Cla I–BspD I 2526
Xmn I 2645
Dsa I 2762
Acl I 4633
I
Msl I 2795
Pac I 4131
BsaB I 3973
Ear I 4073
Eco47 III 3038
1
(Met) Thr Met Ile Thr Asn Ser Ser Ser Val Pro Gly Asp Pro Leu Glu Ser Thr Cys Arg His Ala Ser Leu Ala → LacZ´
6230
Ecl136 II
Sac I
6250
Xma I
Sma I
6270
Xba I
Pst I
6290
Hind III
atgaccatgattacgaattCGAGCTCGGTACCCGGGGATCCTCTAGAGTCGACCTGCAGGCATGCAAGCTTGGcact
EcoR I
Kpn I
BamH I
Sal I
Sph I
Acc65 I
Hinc II
Acc I
Ava I/BsoB I
Fig. 4.17 Restriction map of cloning vector M13mp18. Phages M13mp18 and M13mp19 are 7249 bases in length and differ
only in the orientation of the 54-base polylinker that they carry. The map shows the restriction sites of enzymes that cut the
molecule once or twice. The unique sites are shown in bold type.
making detection of recombinants easy. However,
the lac region only contains unique sites for AvaII,
BglI and PvuI and three sites for PvuII, and there are
no sites anywhere on the complete genome for the
commonly used enzymes such as EcoRI or HindIII.
To remedy this defect, Gronenborn and Messing
(1978) used in vitro mutagenesis to change a single
base pair, thereby creating a unique EcoRI site within
the lac fragment. This variant was designated M13
mp2. This phage derivative was further modified to
generate derivatives with polylinkers upstream of
the lac α-fragment (Fig. 4.17). These derivatives
(mp7–mp11, mp18, mp19) are the exact M13
counterparts of the pUC plasmids shown in Fig. 4.9.
POGC05 9/11/2001 10:57 AM Page 64
CHAPTER 5
Cosmids, phasmids and
other advanced vectors
Introduction
In the 1970s, when recombinant DNA technology
was first being developed, only a limited number
of vectors were available and these were based on
either high-copy-number plasmids or phage λ. Later,
phage M13 was developed as a specialist vector to
facilitate DNA sequencing. Gradually, a number of
purpose-built vectors were developed, of which
pBR322 is probably the best example, but the creation of each one was a major task. Over time, a
series of specialist vectors was constructed, each
for a particular purpose. During this period, there
were many arguments about the relative benefits of
plasmid and phage vectors. Today, the molecular
biologist has available an enormous range of vectors
and these are notable for three reasons. First, many
of them combine elements from both plasmids and
phages and are known as phasmids or, if they contain
an M13 ori region, phagemids. Secondly, many different features that facilitate cloning and expression
can be found combined in a single vector. Thirdly,
purified vector DNA plus associated reagents can
be purchased from molecular-biology suppliers.
The hapless scientist who opens a molecular-biology
catalogue is faced with a bewildering selection of
vectors and each vender promotes different ones.
Although the benefits of using each vector may be
clear, the disadvantages are seldom obvious. The
aim of this chapter is to provide the reader with a
detailed explanation of the biological basis for the
different designs of vector.
There are two general uses for cloning vectors:
cloning large pieces of DNA and manipulating
genes. When mapping and sequencing genomes,
the first step is to subdivide the genome into manageable pieces. The larger these pieces, the easier it is
to construct the final picture (see Chapter 7); hence
the need to clone large fragments of DNA. Large
fragments are also needed if it is necessary to ‘walk’
along the genome to isolate a gene, and this topic is
covered in Chapter 6. In many instances, the desired
gene will be relatively easy to isolate and a simpler
cloning vector can be used. Once isolated, the cloned
gene may be expressed as a probe sequence or as a
protein, it may be sequenced or it may be mutated
in vitro. For all these applications, small specialist
vectors are used.
Vectors for cloning large
fragments of DNA
Cosmid vectors
As we have seen, concatemers of unit-length λ DNA
molecules can be efficiently packaged if the cos sites,
substrates for the packaging-dependent cleavage,
are 37–52 kb apart (75–105% the size of λ+ DNA).
In fact, only a small region in the proximity of the
cos site is required for recognition by the packaging
system (Hohn 1975).
Plasmids have been constructed which contain a
fragment of λ DNA including the cos site (Collins &
Brüning 1978, Collins & Hohn 1979, Wahl et al.
1987, Evans et al. 1989). These plasmids have been
termed cosmids and can be used as gene-cloning
vectors in conjunction with the in vitro packaging
system. Figure 5.1 shows a gene-cloning scheme
employing a cosmid. Packaging the cosmid recombinants into phage coats imposes a desirable selection
upon their size. With a cosmid vector of 5 kb, we
demand the insertion of 32– 47 kb of foreign DNA –
much more than a phage-λ vector can accommodate. Note that, after packaging in vitro, the particle
is used to infect a suitable host. The recombinant
cosmid DNA is injected and circularizes like phage
DNA but replicates as a normal plasmid without
the expression of any phage functions. Transformed cells are selected on the basis of a vector drugresistance marker.
POGC05 9/11/2001 10:57 AM Page 65
Cosmids, phasmids and advanced vectors
65
Cosmid
Restriction
endonuclease
target site
ApR
Foreign DNA restriction
fragment
cos
Restriction
endonuclease
DNA ligase (high DNA concentration)
+
cos
+
+
etc.
cos
37–52 kb
Packaging
in vitro
Transducing particle
containing cosmid
recombinant
Absorption + injection
ApR
cos
Select ApR clone
Fig. 5.1 Simple scheme for cloning in a
cosmid vector. (See text for details.)
Cosmids provide an efficient means of cloning
large pieces of foreign DNA. Because of their capacity for large fragments of DNA, cosmids are particularly attractive vectors for constructing libraries
of eukaryotic genome fragments. Partial digestion
with a restriction endonuclease provides suitably
large fragments. However, there is a potential problem associated with the use of partial digests in
DNA circularizes after
injection
this way. This is due to the possibility of two or
more genome fragments joining together in the
ligation reaction, hence creating a clone containing
fragments that were not initially adjacent in the
genome. This would give an incorrect picture of
their chromosomal organization. The problem can
be overcome by size fractionation of the partial
digest.
POGC05 9/11/2001 10:57 AM Page 66
66
CHAPTER 5
BamHI HindIII
PstI
EcoRI
ApR
EcoRI
cos
pJB8
(5.4 kb)
Sal I
BamHI HindIII
Target genomic
DNA
cos
Sal I
(1) HindIII
(2) Phosphatase
H
HO
cos
H
OH
B
S
HO
B
BamHI
H
HO
(1) SalI
(2) Phosphatase
S
OH
cos
BamHI
B
cos
BH
S
OH HO
B B
S
OH
cos
(1) Partial
Sau3A
digestion
(2) Phosphatase
Sau3A
Sau3A
Mix
Ligate
HO
OH
32–47 kb
H
HO
(B)
(B)
cos
S
cos
OH
37–52 kb
Only packageable molecules
Package
in vitro
Even with sized foreign DNA, in practice cosmid
clones may be produced that contain non-contiguous
DNA fragments ligated to form a single insert.
The problem can be solved by dephosphorylating
the foreign DNA fragments so as to prevent their
ligation together. This method is very sensitive to
the exact ratio of target-to-vector DNAs (Collins &
Brüning 1978) because vector-to-vector ligation
can occur. Furthermore, recombinants with a duplicated vector are unstable and break down in the
Fig. 5.2 Cosmid cloning scheme of
Ish-Horowicz and Burke (1981). (a)
Map of cosmid pJB8. (b) Application
to the construction of a genomic library
of fragments obtained by partial
digestion with Sau3A. This restriction
endonuclease has a tetranucleotide
recognition site and generates
fragments with the same cohesive
termini as BamHI (see p. 31).
host by recombination, resulting in the propagation
of a non-recombinant cosmid vector.
Such difficulties have been overcome in a cosmidcloning procedure devised by Ish-Horowicz and
Burke (1981). By appropriate treatment of the cosmid vector pJB8 (Fig. 5.2), left-hand and right-hand
vector ends are purified which are incapable of selfligation but which accept dephosphorylated foreign
DNA. Thus the method eliminates the need to size
the foreign DNA fragments and prevents formation
POGC05 9/11/2001 10:57 AM Page 67
Cosmids, phasmids and advanced vectors
67
ApR
cos
c2XB
(6.8 kb)
KmR
SmaI
cos
rep
BamHI
Target genomic
DNA
Sma
(Blunt)
Bam
cos
Bam
ApR rep
Sma
(Blunt)
cos
(1) Partial Sau3A digestion
(2) Phosphatase
Sau3A
Sau3A
Mix
Ligate
32–45 kb
Fig. 5.3 Cosmid cloning scheme of
Bates and Swift (1983). The cosmid c2XB
contains two cos sites, separated by a site
for the restriction endonuclease SmaI which
creates blunt ends. These blunt ends ligate
only very inefficiently under the conditions
used and effectively prevent the formation
of recombinants containing multiple
copies of the vector.
Sma
(Blunt)
(B)
cos
(B)
R
Ap rep
37–52 kb
cos
Sma
(Blunt)
Only packageable molecules
of clones containing short foreign DNA or multiple
vector sequences.
An alternative solution to these problems has
been devised by Bates and Swift (1983) who have
constructed cosmid c2XB. This cosmid carries a
BamHI insertion site and two cos sites separated by a
blunt-end restriction site (Fig. 5.3). The creation of
these blunt ends, which ligate only very inefficiently
under the conditions used, effectively prevents
vector self-ligation in the ligation reaction.
Modern cosmids of the pWE and sCos series (Wahl
et al. 1987, Evans et al. 1989) contain features such
as: (i) multiple cloning sites (Bates & Swift 1983,
Pirrotta et al. 1983, Breter et al. 1987) for simple
cloning using non-size-selected DNA; (ii) phage
Package
in vitro
promoters flanking the cloning site; and (iii) unique
NotI, SacII or Sfil sites (rare cutters, see Chapter 6)
flanking the cloning site to permit removal of the
insert from the vector as single fragments. Mammalian expression modules encoding dominant
selectable markers (Chapter 10) may also be present,
for gene transfer to mammalian cells if required.
BACs and PACs as alternatives to cosmids
Phage P1 is a temperate bacteriophage which has
been extensively used for genetic analysis of
Escherichia coli because it can mediate generalized
transduction. Sternberg and co-workers have developed a P1 vector system which has a capacity for
POGC05 9/11/2001 10:57 AM Page 68
68
CHAPTER 5
Short arm
pac loxP
Stuffer
+
P1 and plasmid
replicons: kanr, loxP
Long arm
pac loxP
Insert DNA
Ligate
Insert DNA
P1 and plasmid
replicons: kanr, loxP Stuffer
115 kb
Pac extract cleaves at pac
site and inserts DNA into
P1 heads. Tails are attached
P1 heads
and tails
+
Allow phage to adsorb
to host strain and inject
DNA into cre + host cell
loxP
Cre recombinase protein circularizes injected DNA at the loxP sites. DNA-replicates using
plasmid replicon. Plasmid copy number is increased by induction of P1 lytic replicon.
DNA fragments as large as 100 kb (Sternberg 1990,
Pierce et al. 1992). Thus the capacity is about twice
that of cosmid clones but less than that of yeast
artificial chromosome (YAC) clones (see p. 159). The
P1 vector contains a packaging site (pac) which is
necessary for in vitro packaging of recombinant
molecules into phage particles. The vectors contain
two loxP sites. These are the sites recognized by the
phage recombinase, the product of the phage cre
gene, and which lead to circularization of the packaged DNA after it has been injected into an E. coli
host expressing the recombinase (Fig. 5.4). Clones
are maintained in E. coli as low-copy-number plasmids by selection for a vector kanamycin-resistance
marker. A high copy number can be induced by
Fig. 5.4 The phage P1 vector system.
The P1 vector Ad10 (Sternberg 1990)
is digested to generate short and
long vector arms. These are
dephosphorylated to prevent
self-ligation. Size-selected insert DNA
(85–100 kb) is ligated with vector
arms, ready for a two-stage processing
by packaging extracts. First, the
recombinant DNA is cleaved at the pac
site by pacase in the packaging extract.
Then the pacase works in concert with
head/tail extract to insert DNA into
phage heads, pac site first, cleaving off a
headful of DNA at 115 kb. Heads and
tails then unite. The resulting phage
particle can inject recombinant DNA
into host E. coli. The host is cre+. The
cre recombinase acts on loxP sites to
produce a circular plasmid. The plasmid
is maintained at low copy number, but
can be amplified by inducing the P1
lytic operon.
exploitation of the P1 lytic replicon (Sternberg 1990).
This P1 system has been used to construct genomic
libraries of mouse, human, fission yeast and Drosophila DNA (Hoheisel et al. 1993, Hartl et al. 1994).
Shizuya et al. (1992) have developed a bacterial
cloning system for mapping and analysis of complex
genomes. This BAC system (bacterial artificial chromosome) is based on the single-copy sex factor F of E.
coli. This vector (Fig. 5.5) includes the λ cos N and P1
loxP sites, two cloning sites (HindIII and BamHI) and
several G+C restriction enzyme sites (e.g. SfiI, NotI,
etc.) for potential excision of the inserts. The cloning
site is also flanked by T7 and SP6 promoters for
generating RNA probes. This BAC can be transformed into E. coli very efficiently, thus avoiding the
POGC05 9/11/2001 10:57 AM Page 69
Cosmids, phasmids and advanced vectors
HindIII
XmaI/SmaI
NotI/BgII/Sfi I
BamHI
EagI
NotI
cosN
T7
promoter
IoxP
SP6 promoter
pa
rB
R
Cm
A
ori
S
par
repE
Fig. 5.5 Structure of a BAC vector derived from a mini-F
plasmid. The oriS and repE genes mediate the unidirectional
replication of the F factor, while parA and parB maintain the
copy number at a level of one or two per genome. CmR is a
chloramphenicol-resistance marker. cosN and loxP are the
cleavage sites for λ terminase and P1 cre protein, respectively.
HindIII and BamHI are unique cleavage sites for inserting
foreign DNA. (Adapted from Shizuya et al. 1992.)
packaging extracts that are required with the P1
system. BACs are capable of maintaining human
and plant genomic fragments of greater than 300 kb
for over 100 generations with a high degree of
stability (Woo et al. 1994) and have been used to
construct genome libraries with an average insert
size of 125 kb (Wang et al. 1995a). Subsequently,
Ioannou et al. (1994) have developed a P1-derived
artificial chromosome (PAC), by combining features
of both the P1 and the F-factor systems. Such PAC
vectors are able to handle inserts in the 100–300 kb
range.
The first BAC vector, pBAC108L, lacked a selectable marker for recombinants. Thus, clones with
inserts had to be identified by colony hybridization.
While this once was standard practice in gene manipulation work, today it is considered to be inconvenient! Two widely used BAC vectors, pBeloBAC11
and pECBAC1, are derivatives of pBAC108L in
69
which the original cloning site is replaced with a lacZ
gene carrying a multiple cloning site (Kim et al.
1996, Frijters et al. 1997). pBeloBAC11 has two
EcoRI sites, one in the lacZ gene and one in the CMR
gene, wheras pECBAC1 has only the EcoRI site in
the lacZ gene. Further improvements to BACs have
been made by replacing the lacZ gene with the sacB
gene (Hamilton et al. 1996). Insertional inactivation
of sacB permits growth of the host cell on sucrosecontaining media, i.e. positive selection for inserts.
Frengen et al. (1999) have further improved these
BACs by including a site for the insertion of a transposon. This enables genomic inserts to be modified
after cloning in bacteria, a procedure known as
retrofitting. The principal uses of retrofitting are the
simplified introduction of deletions (Chatterjee &
Coren 1997) and the introduction of reporter genes
for use in the original host of the genomic DNA
(Wang et al. 2001).
Choice of vector
The maximum size of insert that the different vectors
will accommodate is shown in Table 5.1. The size of
insert is not the only feature of importance. The
absence of chimeras and deletions is even more
important. In practice, some 50% of YACs show
structural instability of inserts or are chimeras in
which two or more DNA fragments have become
incorporated into one clone. These defective YACs
are unsuitable for use as mapping and sequencing
reagents and a great deal of effort is required to
identify them. Cosmid inserts sometimes contain
Table 5.1 Maximum DNA insert possible with
different cloning vectors. YACs are discussed on
p. 159.
Vector
l phage
l cosmids
P1 phage
PACs
BACs
YACs
Host
Insert size
E. coli
E. coli
E. coli
E. coli
E coli
Saccharomyces cerevisiae
5–25 kb
35–45 kb
70–100 kb
100–300 kb
≤ 300 kb
200–2000 kb
POGC05 9/11/2001 10:57 AM Page 70
70
CHAPTER 5
the same aberrations and the greatest problem with
them arises when the DNA being cloned contains
tandem arrays of repeated sequences. The problem
is particularly acute when the tandem array is several times larger than the allowable size of a cosmid
insert. Potential advantages of the BAC and PAC
systems over YACs include lower levels of chimerism
(Hartl et al. 1994, Sternberg 1994), ease of library
generation and ease of manipulation and isolation of
insert DNA. BAC clones seem to represent human
DNA far more faithfully than their YAC or cosmid
counterparts and appear to be excellent substrates
for shotgun sequence analysis, resulting in accurate
contiguous sequence data (Venter et al. 1996).
Specialist-purpose vectors
Vectors that can be used to make
single-stranded DNA for sequencing
Whenever a new gene is cloned or a novel genetic
construct is made, it is usual practice to sequence all
or part of the chimeric molecule. As will be seen later
(p. 120), the Sanger method of sequencing requires
single-stranded DNA as the starting material.
Originally, single-stranded DNA was obtained by
cloning the sequence of interest in an M13 vector
(see p. 60). Today, it is more usual to clone the
sequence into a pUC-based phagemid vector which
contains the M13 ori region as well as the pUC
(Col E1) origin of replication. Such vectors normally replicate inside the cell as double-stranded
molecules. Single-stranded DNA for sequencing can
be produced by infecting cultures with a helper
phage such as M13K07. This helper phage has the
origin of replication of P15A and a kanamycin-
-35 Region
resistance gene inserted into the M13 ori region and
carries a mutation in the gII gene (Vieira & Messing
1987). M13KO7 can replicate on its own. However,
in the presence of a phagemid bearing a wild-type
origin of replication, single-stranded phagemid is
packaged preferentially and secreted into the culture medium. DNA purified from the phagemids can
be used directly for sequencing.
Expression vectors
Expression vectors are required if one wants to prepare RNA probes from the cloned gene or to purify
large amounts of the gene product. In either case,
transcription of the cloned gene is required. Although
it is possible to have the cloned gene under the control of its own promoter, it is more usual to utilize a
promoter specific to the vector. Such vector-carried
promoters have been optimized for binding of the E.
coli RNA polymerase and many of them can be regulated easily by changes in the growth conditions of
the host cell.
E. coli RNA polymerase is a multi-subunit enzyme.
The core enzyme consists of two identical α subunits
and one each of the β and β′ subunits. The core
enzyme is not active unless an additional subunit,
the σ factor, is present. RNA polymerase recognizes
different types of promoters depending on which
type of σ factor is attached. The most common promoters are those recognized by the RNA polymerase
with σ70. A large number of σ70 promoters from E. coli
have been analysed and a compilation of over 300 of
them can be found in Lisser and Margalit (1993). A
comparison of these promoters has led to the formulation of a consensus sequence (Fig. 5.6). If the transcription start point is assigned the position +1 then
-10 Region
1 2 3 4 5 6 7 8 9
1011121314151617
CONSENSUS
• • • T TGACA • • • • • • • • •
lac
GGC T T T A C A C T T T A T G C T T C C GG C T C G T A T A T T G T
• • • • • • • •TATAAT• •
trp
C TGT TGACA A T T A A T CA T
λPL
GTGT TGACA T A A A T ACCA
C T GG C GG T G A T A C TGA
rec A
CAC T TGA T AC TGT A TGA A
GC A T A C A G T A T A A T TG
tacI
C TGT TGACA A T T A A T CA T
CGGC T CG T A T A A TGT
tacII
C TGT TGACA A T T A A T CA T
CG A A C T A G T T T A A TGT
CG A A C T A G T T A A C T AG
Fig. 5.6 The base sequence of the
−10 and −35 regions of four natural
promoters, two hybrid promoters and
the consensus promoter.
POGC05 9/11/2001 10:57 AM Page 71
Cosmids, phasmids and advanced vectors
this consensus sequence consists of the −35 region
(5′-TTGACA-) and the −10 region, or Pribnow box
(5′-TATAAT). RNA polymerase must bind to both
sequences to initiate transcription. The strength of a
promoter, i.e. how many RNA copies are synthesized per unit time per enzyme molecule, depends on
how close its sequence is to the consensus. While
the −35 and −10 regions are the sites of nearly all
mutations affecting promoter strength, other bases
flanking these regions can affect promoter activity
(Hawley & McClure 1983, Dueschle et al. 1986,
Keilty & Rosenberg 1987). The distance between the
−35 and −10 regions is also important. In all cases
examined, the promoter was weaker when the
spacing was increased or decreased from 17 bp.
Upstream (UP) elements located 5′ of the −35 hexamer in certain bacterial promoters are A+T-rich
sequences that increase transcription by interacting
with the α subunit of RNA polymerase. Gourse et al.
(1998) have identified UP sequences conferring
increased activity to the rrn core promoter. The best
UP sequence was portable and increased heterologous protein expression from the lac promoter by a
factor of 100.
Once RNA polymerase has initiated transcription
at a promoter, it will polymerize ribonucleotides
until it encounters a transcription-termination
site in the DNA. Bacterial DNA has two types of
transcription-termination site: factor-independent
and factor-dependent. As their names imply, these
types are distinguished by whether they work with
just RNA polymerase and DNA alone or need other
factors before they can terminate transcription.
The factor-independent transcription terminators
are easy to recognize because they have similar
sequences: an inverted repeat followed by a string of
A residues (Fig. 5.7). Transcription is terminated
in the string of A residues, resulting in a string of
GC-rich inverted repeat
5’ - G T C A A A A G C C T C C G G T C G G A G G C T T T T G A C T
C A G T T T T C G G A G G C C A G C C T C C G A A A A C T G A- 5’
Run of
A residues
Fig. 5.7 Structure of a factor-independent transcriptional
terminator.
71
U residues at the 3′ end of the mRNA. The factordependent transcription terminators have very little
sequence in common with each other. Rather,
termination involves interaction with one of the
three known E. coli termination factors, Rho (ρ),
Tau (τ) and NusA. Most expression vectors incorporate a factor-independent termination sequence
downstream from the site of insertion of the cloned
gene.
Vectors for making RNA probes
Although single-stranded DNA can be used as a
sequence probe in hybridization experiments, RNA
probes are preferred. The reasons for this are that the
rate of hybridization and the stability are far greater
for RNA–DNA hybrids compared with DNA–DNA
hybrids. To make an RNA probe, the relevant gene
sequence is cloned in a plasmid vector such that it
is under the control of a phage promoter. After
purification, the plasmid is linearized with a suitable
restriction enzyme and then incubated with the
phage RNA polymerase and the four ribonucleoside
triphosphates (Fig. 5.8). No transcription terminator is required because the RNA polymerase will fall
off the end of the linearized plasmid.
There are three reasons for using a phage promoter. First, such promoters are very strong, enabling
large amounts of RNA to be made in vitro. Secondly,
the phage promoter is not recognized by the E. coli
RNA polymerase and so no transcription will occur
inside the cell. This minimizes any selection of variant inserts. Thirdly, the RNA polymerases encoded
by phages such as SP6, T7 and T3 are much simpler
molecules to handle than the E. coli enzyme, since
the active enzyme is a single polypeptide.
If it is planned to probe RNA or single-stranded
DNA sequences, then it is essential to prepare RNA
probes corresponding to both strands of the insert.
One way of doing this is to have two different clones
corresponding to the two orientations of the insert.
An alternative method is to use a cloning vector in
which the insert is placed between two different,
opposing phage promoters (e.g. T7/T3 or T7/SP6)
that flank a multiple cloning sequence (see Fig. 5.5).
Since each of the two promoters is recognized by a
different RNA polymerase, the direction of transcription is determined by which polymerase is used.
POGC05 9/11/2001 10:57 AM Page 72
72
CHAPTER 5
Probe
sequence
SP6 promoter
Insert probe
Vector
sequence
Linearize
plasmid
Add SP6 RNA polymerase.
Unlabelled CTP, ATP, GTP.
Labelled UTP
High specific activity
RNA probes
(a)
pUC
origin
M13
origin
pLITMUSTM
2.8 kb
ampr
lacZ’
LITMUS 28
LITMUS 29
SnaBI
SpeI
SnaBI
SpeI
T7
T7
Fig. 5.8 Method for preparing RNA
probes from a cloned DNA molecule
using a phage SP6 promoter and SP6
RNA polymerase.
LITMUS 38
SnaBI
SpeI
T7
LITMUS 39
SnaBI
SpeI
T7
Bgl II
NsiI - Ppu 10I
BssHII
BsiWI
XhoI
EcoRI
PstI
EcoRV
BamHI
HindIII
NcoI
AatII
AgeI
XbaI
AvrII
SacI
KpnI - Acc 65I
StuI
KpnI-Acc65I
SacI
AvrII
XbaI
AgeI
AatII
NcoI
HindIII
BamHI
EcoRV
PstI
EcoRI
XhoI
BsiW I
BssHII
NsiI - Ppu10I
Bgl II
StuI
ApaI-Bsp120I
MfeI
NgoMI
KasI
HindIII
PstI
EcoRV
BamHI
EcoRI
NheI
EagI
MluI
BspE II
Bsr GI
SphI
SalI
StuI
SalI
SphI
Bsr GII
BspE I
MluI
EagI
NheI
EcoRI
BamHI
EcoRV
PstI
HindIII
KasI
NgoMI
MfeI
Apa-Bsp120I
StuI
Afl II
Afl II
Afl II
Afl II
T7
T7
T7
T7
Fig. 5.9 Structure and use of the LITMUS vectors for making RNA probes. (a) Structure of the LITMUS vectors showing the
orientation and restriction sites of the four polylinkers.
POGC05 9/11/2001 10:57 AM Page 73
Cosmids, phasmids and advanced vectors
(b)
RNA transcript of top
strand of insert
5’
3’
Table 5.2 Factors affecting the expression of
cloned genes.
Factor
T7 RNA Polymerase,
rNTP’s
AfI II
Spe I
AfI II
T7
Linearize with
Afl II
73
Promoter strength
Transcriptional termination
Plasmid copy number
Plasmid stability
Host-cell physiology
Translational initiation sequences
Codon choice
mRNA structure
Text
Page 74
Page 71
Page 45, Chapter 4
Page 47, Chapter 4
Chapters 4 & 5
Box 5.1, page 77
Box 5.1, page 77
Box 5.1, page 77
Spe I
T7
Cloned
insert
LITMUS
T7
Afl II
Linearize with
SpeI
SpeI
SpeI
Afl II
T7
T7 RNA Polymerase,
rNTP’s
3’
5’
RNA transcript of
bottom strand of insert
Fig. 5.9 (cont’d ) (b) Method of using the LITMUS vectors to
selectively synthesize RNA probes from each strand of a
cloned insert. (Figure reproduced courtesy of New England
Biolabs.)
A further improvement has been introduced
by Evans et al. (1995). In their LITMUS vectors,
the polylinker regions are flanked by two modified
T7 RNA polymerase promoters. Each contains a
unique restriction site (SpeI or AflII) that has been
engineered into the T7 promoter consensus sequence
such that cleavage with the corresponding endonuclease inactivates that promoter. Both promoters
are active despite the presence of engineered sites.
Selective unidirectional transcription is achieved by
simply inactivating the other promoter by digestion
with SpeI or AflII prior to in vitro transcription
(Fig. 5.9). Since efficient labelling of RNA probes
demands that the template be linearized prior to
transcription, at a site downstream from the insert,
cutting at the site within the undesired promoter
performs both functions in one step. Should the
cloned insert contain either an SpeI or an AflII site,
the unwanted promoter can be inactivated by cutting at one of the unique sites within the polylinker.
Vectors for maximizing protein synthesis
Provided that a cloned gene is preceded by a promoter recognized by the host cell, then there is a
high probability that there will be detectable synthesis of the cloned gene product. However, much
of the interest in the application of recombinant
DNA technology is the possibility of facile synthesis
of large quantities of protein, either to study its
properties or because it has commercial value. In
such instances, detectable synthesis is not sufficient:
rather, it must be maximized. The factors affecting
the level of expression of a cloned gene are shown
in Table 5.2 and are reviewed by Baneyx (1999).
Of these factors, only promoter strength is considered here.
POGC05 9/11/2001 10:57 AM Page 74
74
CHAPTER 5
Promoter
l PL
trc
T7
Uninduced level of CAT
Induced level of CAT
Ratio
0.0275
1.10
1.14
28.18
5.15
15.40
1025
4.7
13.5
When maximizing gene expression it is not enough
to select the strongest promoter possible: the effects
of overexpression on the host cell also need to be considered. Many gene products can be toxic to the host
cell even when synthesized in small amounts.
Examples include surface structural proteins (Beck
& Bremer 1980), proteins, such as the PolA gene
product, that regulate basic cellular metabolism
(Murray & Kelley 1979), the cystic fibrosis transmembrane conductance regulator (Gregory et al.
1990) and lentivirus envelope sequences (Cunningham et al. 1993). If such cloned genes are allowed
to be expressed there will be a rapid selection for
mutants that no longer synthesize the toxic protein.
Even when overexpression of a protein is not toxic to
the host cell, high-level synthesis exerts a metabolic
drain on the cell. This leads to slower growth and
hence in culture there is selection for variants with
lower or no expression of the cloned gene because
these will grow faster. To minimize the problems
associated with high-level expression, it is usual to
use a vector in which the cloned gene is under the
control of a regulated promoter.
Many different vectors have been constructed for
regulated expression of gene inserts but most of
those in current use contain one of the following
controllable promoters: λ PL , T7, trc (tac) or BAD.
Table 5.3 shows the different levels of expression that
can be achieved when the gene for chloramphenicol
transacetylase (CAT) is placed under the control of
three of these promoters.
The trc and tac promoters are hybrid promoters
derived from the lac and trp promoters (Brosius
1984). They are stronger than either of the two
parental promoters because their sequences are
more like the consensus sequence. Like lac, the trc
and tac promoters are inducibile by lactose and
isopropyl-β-d-thiogalactoside (IPTG). Vectors using
Table 5.3 Control of expression of
chloramphenicol acetyltransferase
(CAT) in E. coli by three different
promoters. The levels of CAT are
expressed as µg/mg total protein.
these promoters also carry the lacO operator and the
lacI gene, which encodes the repressor.
The pET vectors are a family of expression vectors
that utilize phage T7 promoters to regulate synthesis
of cloned gene products (Studier et al. 1990). The
general strategy for using a pET vector is shown in
Fig. 5.10. To provide a source of phage-T7 RNA
polymerase, E. coli strains that contain gene 1 of the
phage have been constructed. This gene is cloned
downstream of the lac promoter, in the chromosome, so that the phage polymerase will only be
synthesized following IPTG induction. The newly
synthesized T7 RNA polymerase will then transcribe
the foreign gene in the pET plasmid. If the protein
product of the cloned gene is toxic, it is possible to
minimize the uninduced level of T7 RNA polymerase. First, a plasmid compatible with pET vectors
is selected and the T7 lysS gene is cloned in it. When
introduced into a host cell carrying a pET plasmid,
the lysS gene will bind any residual T7 RNA polymerase (Studier 1991, Zhang & Studier 1997). Also,
if a lac operator is placed between the T7 promoter
and the cloned gene, this will further reduce transcription of the insert in the absence of IPTG
(Dubendorff & Studier 1991). Improvements in the
yield of heterologous proteins can sometimes be
achieved by use of selected host cells (Miroux &
Walker 1996).
The λ PL promoter system combines very tight
transcriptional control with high levels of gene
expression. This is achieved by putting the cloned
gene under the control of the PL promoter carried on
a vector, while the PL promoter is controlled by a cI
repressor gene in the E. coli host. This cI gene is itself
under the control of the tryptophan (trp) promoter
(Fig. 5.11). In the absence of exogenous tryptophan,
the cI gene is transcribed and the cI repressor binds
to the PL promoter, preventing expression of the
POGC05 9/11/2001 10:57 AM Page 75
Cosmids, phasmids and advanced vectors
IPTG induction
75
IPTG Induction
Host cell
E.coli RNA
polymerase
T7 RNA
polymerase
T7 gene 1
Target gene
T7 RNA polymerase
IacO
Iac promoter
IacO
T7 promoter
Inactive
DE3
lac
repressor
Iac
repressor
Iacl gene
pET
lacl gene
T7 lysozyme
T7 lysozyme
gene
pLysS
or E
E.coli genome
Fig. 5.10 Strategy for regulating the expression of genes cloned into a pET vector. The gene for T7 RNA polymerase (gene 1) is
inserted into the chromosome of E. coli and transcribed from the lac promoter; therefore, it will be expressed only if the inducer
IPTG is added. The T7 RNA polymerase will then transcribe the gene cloned into the pET vector. If the protein product of the
cloned gene is toxic, it may be necessary to further reduce the transcription of the cloned gene before induction. The T7 lysozyme
encoded by a compatible plasmid, pLysS, will bind to any residual T7 RNA polymerase made in the absence of induction and
inactivate it. Also, the presence of lac operators between the T7 promoter and the cloned gene will further reduce transcription
of the cloned gene in the absence of the inducer IPTG. (Reprinted with permission from the Novagen Catalog, p. 24, Novagen,
Madison, Wisconsin, 1995.)
Minus tryptophan
Plus tryptophan
Tryptophan
Transcription
PO
Ptrp
No transcription
λcI repressor
PO
cI repressor
ATG
Bacterial
chromosome
Transcription
No transcription
PL
λcI repressor
trp repressor
Bacterial
chromosome
PO
Ptrp
PO
Gene of interest
Vector
PL
ATG
Gene of interest
Vector
Fig. 5.11 Control of cloned gene expression using the λcI promoter. See text for details. (Diagram reproduced courtesy of
In Vitrogen Corporation.)
cloned gene. Upon addition of tryptophan, the trp
repressor binds to the cI gene, preventing synthesis
of the cI repressor. In the absence of cI repressor,
there is a high level of expression from the very
strong PL promoter.
The pBAD vectors, like the ones based on PL
promoter, offer extremely tight control of expression
of cloned genes (Guzman et al. 1995). The pBAD
vectors carry the promoter of the araBAD (arabinose) operon and the gene encoding the positive and
POGC05 9/11/2001 10:57 AM Page 76
76
CHAPTER 5
(a)
O2
C
N
AraC dimer
N
C
Pc
No transcription
I1
I2
pBAD
the glucose and arabinose content of the growth
medium (Fig. 5.12). According to Guzman et al.
(1995), the pBAD vectors permit fine-tuning of gene
expression. All that is required is to change the sugar
composition of the medium. However, this is disputed
by others (Siegele & Hu 1997, Hashemzadeh-Bonehi
et al. 1998).
Many of the vectors designed for high-level expression also contain translation-initiation signals
optimized for E. coli expression (see Box 5.1).
+ arabinose
Vectors to facilitate protein purification
N
N
Transcription
Pc
CAP
C
C
I1
I2
pBAD
(b)
Fig. 5.12 Regulation of the pBAD promoter. (a) The
conformational changes that take place on addition of
arabinose. (b) Western blot showing the increase in synthesis
of a cloned gene product when different levels of arabinose are
added to a culture of the host cell.
negative regulator of this promoter, araC. AraC is a
transcriptional regulator that forms a complex with
-arabinose. In the absence of arabinose, AraC binds
to the O2 and I1 half-sites of the araBAD operon,
forming a 210 bp DNA loop and thereby blocking
transcription (Fig. 5.12). As arabinose is added to
the growth medium, it binds to AraC, thereby releasing the O2 site. This in turn causes AraC to bind to
the I2 site adjacent to the I1 site. This releases the
DNA loop and allows transcription to begin. Binding
of AraC to I1 and I2 is activated in the presence of
cAMP activator protein (CAP) plus cAMP. If glucose
is added to the growth medium, this will lead to a
repression of cAMP synthesis, thereby decreasing
expression from the araBAD promoter. Thus one can
titrate the level of cloned gene product by varying
Many cloning vectors have been engineered so that
the protein being expressed will be fused to another
protein, called a tag, which can be used to facilitate
protein purification. Examples of tags include glutathione-S-transferase, the MalE (maltose-binding)
protein and multiple histidine residues, which can
easily be purified by affinity chromatography. The
tag vectors are usually constructed so that the
coding sequence for an amino acid sequence cleaved
by a specific protease is inserted between the coding
sequence for the tag and the gene being expressed.
After purification, the tag protein can be cleaved off
with the specific protease to leave a normal or nearly
normal protein. It is also possible to include in the
tag a protein sequence that can be assayed easily.
This permits assay of the cloned gene product when
its activity is not known or when the usual assay is
inconvenient. Three different examples of tags are
given below. The reader requiring a more detailed
insight should consult the review by LaVallie and
McCoy (1995).
To use a polyhistidine fusion for purification, the
gene of interest is first engineered into a vector in
which there is a polylinker downstream of six histidine residues and a proteolytic cleavage site. In the
example shown in Fig. 5.13, the cleavage site is that
for enterokinase. After induction of synthesis of the
fusion protein, the cells are lysed and the viscosity
of the lysate is reduced by nuclease treatment.
The lysate is then applied to a column containing
immobilized divalent nickel, which selectively binds
the polyhistidine tag. After washing away any
contaminating proteins, the fusion protein is eluted
from the column and treated with enterokinase to
release the cloned gene product.
POGC05 9/11/2001 10:57 AM Page 77
Cosmids, phasmids and advanced vectors
Box 5.1 Optimizing translation
High-level expression of a cloned gene requires more
than a strong promoter. The mRNA produced during
transcription needs to be effectively translated into
protein. Although many factors can influence the rate
of translation, the most important is the interaction
of the ribosome with the bases immediately upstream
from the initiation codon of the gene. In bacteria,
a key sequence is the ribosome-binding site or
Shine–Dalgarno (S-D) sequence. The degree of
complementarity of this sequence with the 16S rRNA
can affect the rate of translation (De Boer & Hui
1990). Maximum complementarity occurs with the
sequence 5′-UAAGGAGG-3′ (Ringquist et al. 1992).
The spacing between the S-D sequence and the
initiation codon is also important. Usually there are
five to 10 bases, with eight being optimal.
Decreasing the distance below 4 bp or increasing it
beyond 14 bp can reduce translation by several
orders of magnitude.
Translation is affected by the sequence of bases
that follow the S-D site (De Boer et al. 1983b). The
presence of four A residues or four T residues in this
position gave the highest translational efficiency.
Translational efficiency was 50% or 25% of maximum
when the region contained, respectively, four C
residues or four G residues.
The composition of the triplet immediately
preceding the AUG start codon also affects
the efficiency of translation. For translation of
b-galactosidase mRNA, the most favourable
combinations of bases in this triplet are UAU and
CUU. If UUC, UCA or AGG replaced UAU or CUU, the
level of expression was 20-fold less (Hui et al. 1984).
The codon composition following the AUG
start codon can also affect the rate of translation.
For example, a change in the third position of
the fourth codon of a human g-interferon gene
resulted in a 30-fold change in the level of expression
(De Boer & Hui 1990). Also, there is a strong bias in
the second codon of many natural mRNAs, which is
quite different from the general bias in codon usage.
Highly expressed genes have AAA (Lys) or GCU (Ala)
as the second codon. DevIin et al. (1988) changed all
the G and C nucleotides for the first four codons of a
granulocyte colony-stimulating factor gene and
expression increased from undetectable to 17% of
total cell protein.
Sequences upstream from the S-D site can affect
the efficiency of translation of certain genes. In the
E. coli rnd gene there is a run of eight uracil residues.
Changing two to five of these residues has no effect
on mRNA levels but reduces translation by up to 95%
(Zhang & Deutscher 1992). Etchegaray and Inouye
(1999) have identified an element downstream of
the initiation codon, the downstream box, which
facilitates formation of the translation-initiation
complex. The sequence of the 3′ untranslated region
of the mRNA can also be important. If this region is
complementary to sequences within the gene, hairpin
loops can form and hinder ribosome movement along
the messenger.
The genetic code is degenerate, and hence for
most of the amino acids there is more than one
codon. However, in all genes, whatever their origin,
the selection of synonymous codons is distinctly
non-random (for reviews, see Kurland 1987, Ernst
1988 and McPherson 1988). The bias in codon
usage has two components: correlation with tRNA
availability in the cell, and non-random choices
between pyrimidine-ending codons. Ikemura (1981a)
measured the relative abundance of the 26 known
tRNAs of E. coli and found a strong positive correlation
between tRNA abundance and codon choice. Later,
Ikemura (1981b) noted that the most highly expressed
genes in E. coli contain mostly those codons
corresponding to major tRNAs but few codons of
minor tRNAs. In contrast, genes that are expressed
less well use more suboptimal codons. Forman et al.
(1998) noted significant misincorporation of lysine,
in place of arginine, when the rare AGA codon was
included in a gene overexpressed in E. coli. It should
be noted that the bias in codon usage even extends
to the stop codons (Sharp & Bulmer 1988). UAA is
favoured in genes expressed at high levels, whereas
UAG and UGA are used more frequently in genes
expressed at a lower level.
For a review of translation the reader should
consult Kozak (1999).
77
POGC05 9/11/2001 10:57 AM Page 78
CHAPTER 5
XhoI
SacI*
BglII
PstI
PvuII*
KpnI
EcoRI
SfuI
HindIII
78
EK MCS
ATG 6xHis Epitope Site
term
Ampicillin
araC
PBAD
pBAD/His
A,B,C
4.1 kb
ColE1
*Frame-dependent variations
COMMON SEQUENCE
VARIABLE SEQUENCE
GAC GAT GAC GAT AAG GAT
Asp
Asp
Asp
Asp Lys
Asp
GAC GAT GAC GAT AAG GAT
Asp
Asp
Asp
Asp Lys
Asp
GAC GAT GAC GAT AAG GAT
Asp
Asp
Asp
Asp Lys
Asp
CCG AGC TCG AGA TCT GCA GCT
Pro
Ser
Ser
Ars
Ser
Ala
Ala
CGA TGG GGA TCC GAG CTC GAG ATC TGC
Arg
Trp
Gly
Ser
Glu
Leu
Glu
Ile
Cys
CGA TGG ATC CGA CCT CGA GAT CTG CAG
Arg
Trp
Ile
Arg
Pro
Arg Asp
Leu
Gln
Enterokinase
cleavage
Fig. 5.13 Structure of a vector (pBAD/His, In Vitrogen Corporation) designed for the expression of a cloned gene as a fusion
protein containing a polyhistidine sequence. Three different variants (A, B, C) allow the insert to be placed in each of the three
translational reading frames. The sequence shaded pink shows the base sequence which is altered in each of the three vectors.
The lightly-shaded box (AGATCT) is the Bgl II site of the polylinker. Note that the initial A residue of the restriction site is at a
different point in the triplet codon in each of the three sequences.
For the cloned gene to be expressed correctly, it
has to be in the correct translational reading frame.
This is achieved by having three different vectors,
each with a polylinker in a different reading frame
(see Fig. 5.13). Enterokinase recognizes the sequence
(Asp)4Lys and cleaves immediately after the lysine
residue. Therefore, after enterokinase cleavage, the
cloned gene protein will have a few extra amino
acids at the N terminus. If desired, the cleavage
site and polyhistidines can be synthesized at the C
terminus. If the cloned gene product itself contains
an enterokinase cleavage site, then an alternative
protease, such as thrombin or factor Xa, with a
different cleavage site can be used.
To facilitate assay of the fusion proteins, short
antibody recognition sequences can be incorporated
into the tag between the affinity label and the protease cleavage site. Some examples of recognizable
POGC05 9/11/2001 10:57 AM Page 79
Cosmids, phasmids and advanced vectors
Table 5.4 Peptide epitopes, and the
antibodies that recognize them, for use
in assaying fusion proteins.
Peptide sequence
Antibody recognition
-Glu-Gln-Lys-Leu-Ile Ser-GIu-Glu-Asp-Leu-His-His-His-His-His-His-COOH
-Gly-Lys-Pro-Ile-Pro-Asn-Pro-Leu-Leu-Gly-LeuAsp-Ser-Thr-
epitopes are given in Table 5.4. These antibodies can
be used to detect, by western blotting, fusion proteins carrying the appropriate epitope. Note that a
polyhistidine tag at the C terminus can function for
both assay and purification.
Biotin is an essential cofactor for a number of
carboxylases important in cell metabolism. The
biotin in these enzyme complexes is covalently
attached at a specific lysine residue of the biotin
carboxylase carrier protein. Fusions made to a
segment of the carrier protein are recognized in E.
coli by biotin ligase, the product of the birA gene, and
biotin is covalently attached in an ATP-dependent
reaction. The expressed fusion protein can be
purified using streptavidin affinity chromatography
(Fig. 5.14). E. coli expresses a single endogenous
biotinylated protein, but it does not bind to streptavidin in its native configuration, making the affinity
purification highly specific for the recombinant
fusion protein. The presence of biotin on the fusion
protein has an additional advantage: its presence can
be detected with enzymes coupled to streptavidin.
The affinity purification systems described above
suffer from the disadvantage that a protease is
required to separate the target protein from the
affinity tag. Also, the protease has to be separated
from the protein of interest. Chong et al. (1997,
1998) have described a unique purification system
that has neither of these disadvantages. The system
utilizes a protein splicing element, an intein, from
the Saccharomyces cerevisiae VMA1 gene (see Box 5.2).
The intein is modified such that it undergoes a
self-cleavage reaction at its N terminus at low
temperatures in the presence of thiols, such as
cysteine, dithiothreitol or β-mercaptoethanol. The
gene encoding the target protein is inserted into a
multiple cloning site (MCS) of a vector to create a
fusion between the C terminus of the target gene and
79
Anti-myc antibody
Anti-His (C-terminal) antibody
Anti-V5 antibody
the N terminus of the gene encoding the intein. DNA
encoding a small (5 kDa) chitin-binding domain
from Bacillus circulans was added to the C terminus
of the intein for affinity purification (Fig. 5.15).
The above construct is placed under the control of
an IPTG-inducible T7 promoter. When crude extracts
from induced cells are passed through a chitin column, the fusion protein binds and all contaminating
proteins are washed through. The fusion is then
induced to undergo intein-mediated self-cleavage on
the column by incubation with a thiol. This releases
the target protein, while the intein chitin-binding
domain remains bound to the column.
Vectors to promote solubilization of
expressed proteins
One of the problems associated with the overproduction of proteins in E. coli is the sequestration of the
product into insoluble aggregates or ‘inclusion bodies’ (Fig. 5.16). They were first reported in strains
overproducing insulin A and B chains (Williams
et al. 1982). At first, their formation was thought to
be restricted to the overexpression of heterologous
proteins in E. coli, but they can form in the presence
of high levels of normal E. coli proteins, e.g. subunits
of RNA polymerase (Gribskov & Burgess 1983).
Two parameters that can be manipulated to reduce inclusion-body formation are temperature and
growth rate. There are a number of reports which
show that lowering the temperature of growth
increases the yield of correctly folded, soluble protein
(Schein & Noteborn 1988, Takagi et al. 1988,
Schein 1991). Media compositions and pH values
that reduce the growth rate also reduce inclusionbody formation. Renaturation of misfolded proteins
can sometimes be achieved following solubilization
in guanidinium hydrochloride (Lilie et al. 1998).
POGC05 9/11/2001 10:57 AM Page 80
80
CHAPTER 5
ce Factor Xa
sequen
MC
tag
Sc
lon
ed
ge
n
e
cP
ta
ori
AmpR
Express fusion protein in E. coli
Lyse cells
biotin
tag
Xa
gene product
Affinity chromatography with streptavidin column
Streptavidin
biotin
tag
Streptavidin
Xa
gene product
Elute by adding excess biotin
biotin
tag
Xa
gene product
Cleave fusion protein with factor Xa
biotin
tag
gene product
Fig. 5.14 Purification of a cloned gene
product synthesized as a fusion to the
biotin carboxylase carrier protein (tag).
See text for details.
POGC05 9/11/2001 10:57 AM Page 81
Cosmids, phasmids and advanced vectors
81
Box 5.2 Inteins, exeins and protein splicing
Protein splicing is defined as the excision of an
intervening protein sequence (the intein) from a
protein precursor. Splicing involves ligation of the
flanking protein fragments (the exteins) to form a
mature extein protein and the free intein. Protein
splicing results in a native peptide bond between the
ligated exteins and this differentiates it from other
forms of autoproteolysis.
N-extein
Intein
C-extein
Precursor protein
Splicing
N-extein
C-extein
Intein
Mature extein protein
Sequence comparison and structural analysis have
indicated that the residues responsible for splicing
are ~100 amino acids at the N terminus of the intein
and ~50 amino acids at the C terminus. These two
splicing regions are separated by a linker or a gene
encoding a homing endonuclease. If present in a cell,
N-extein
N-terminal
splicing region
Three ‘genetic’ methods of preventing inclusionbody formation have been described. In the first of
these, the host cell is engineered to overproduce a
chaperon (e.g. DnaK, GroEL or GroES proteins) in
addition to the protein of interest (Van Dyk et al.
1989, Blum et al. 1992, Thomas et al. 1997).
Castanie et al. (1997) have developed a series of
vectors which are compatible with pBR322-type
plasmids and which encode the overproduction of
chaperons. These vectors can be used to test the
effect of chaperons on the solubilization of heterologous gene products. Even with excess chaperon
there is no guarantee of proper folding. The second
method involves making minor changes to the
amino acid sequence of the target protein. For
example, cysteine-to-serine changes in fibroblast
growth factor minimized inclusion-body formation
(Rinas et al. 1992). The third method is derived from
the observation that many proteins produced as
the homing endonuclease makes a double-stranded
break in the DNA at or near the insertion site (the
home) of the intein encoding it. This endonuclease
activity initiates the movement of the intein into
another allele of the same gene if that allele lacks
the intein.
Linker or homing
endonuclease gene
C-terminal
splicing region
C-extein
insoluble aggregates in their native state are
synthesized in soluble form as thioredoxin fusion
proteins (LaVallie et al. 1993). More recently, Davis
et al. (1999) have shown that the NusA and GrpE
proteins, as well as bacterioferritin, are even better
than thioredoxin at solubilizing proteins expressed
at a high level. Kapust and Waugh (1999) have
reported that the maltose-binding protein is also
much better than thioredoxin.
Building on the work of LaVallie et al. (1993), a
series of vectors has been developed in which the
gene of interest is cloned into an MCS and the gene
product is produced as a thioredoxin fusion protein
with an enterokinase cleavage site at the fusion
point. After synthesis, the fusion protein is released
from the producing cells by osmotic shock and
purified. The desired protein is then released by
enterokinase cleavage. To simplify the purification
of thioredoxin fusion proteins, Lu et al. (1996)
POGC05 9/11/2001 10:57 AM Page 82
82
CHAPTER 5
Cleave
Target Gene
CBD
T7 promoter
MCS
Cloning and
expression
Target Protein
Load F.T.
Intein
Strip with
Elute SDS
Wash +DTT
–IPTG +IPTG
CBD
4°C
o.n.
Intein
Intein
Load and
wash
Chitin
Inducible cleavage
+DTT at 4°C
kDa
212
158
116
97.2
66.4
55.6
Precursor
Intein – CBD
MBP
42.7
Elute
Target Protein
36.5
26.6
20.0
1
2
3
4
5
6
7
8
9 10
Lane 1: Protein Marker.
Lane 2: Crude extract from uninduced cells.
Lane 3: Crude extract from cells, induced at 15°C for 16 hours.
Lane 4: Clarified crude extract from induced cells.
Lane 5: Chitin column flow through (F.T.).
Lane 6: Chitin column wash.
Lane 7: Quick DTT wash to distribute DTT evenly throughout the chitin column.
Lanes 8-9: Fraction of eluted MBP after stopping column flow and inducing a self-cleavage reaction at 4°C overnight.
Lane 10: SDS stripping of remaining proteins bound to chitin column (mostly the cleaved intein-CBD fusion).
Fig. 5.15 Purification of a cloned gene product synthesized as a fusion with an intein protein. (Figure reproduced courtesy of New
England Biolabs.)
systematically mutated a cluster of surface amino
acid residues. Residues 30 and 62 were converted
to histidine and the modified (‘histidine patch’)
thioredoxin could now be purified by affinity chromatography on immobilized divalent nickel. An
alternative purification method was developed by
Smith et al. (1998). They synthesized a gene in
which a short biotinylation peptide is fused to the N
terminus of the thioredoxin gene to generate a new
protein called BIOTRX. They constructed a vector
carrying the BIOTRX gene, with an MCS at the C terminus, and the birA gene. After cloning a gene in
the MCS, a fused protein is produced which can be
purified by affinity chromatography on streptavidin
columns.
An alternative way of keeping recombinant proteins soluble is to export them to the periplasmic
space (see next section). However, even here they
may still be insoluble. Barth et al. (2000) solved this
problem by growing the producing bacteria under
osmotic stress (4% NaCl plus 0.5 mol/l sorbitol) in
the presence of compatible solutes. Compatible
solutes are low-molecular-weight osmolytes, such
as glycine betaine, that occur naturally in halophilic
bacteria and are known to protect proteins at high
salt concentrations. Adding glycine betaine for the
cultivation of E. coli under osmotic stress not only
allowed the bacteria to grow under these otherwise
inhibitory conditions but also produced a periplasmic environment for the generation of correctly
folded recombinant proteins.
Vectors to promote protein export
Gram-negative bacteria such as E. coli have a complex wall–membrane structure comprising an inner,
cytoplasmic membrane separated from an outer
membrane by a cell wall and periplasmic space.
POGC05 9/11/2001 10:57 AM Page 83
Cosmids, phasmids and advanced vectors
83
Fig. 5.16 Inclusions of Trp polypeptide–proinsulin fusion protein in E. coli. (Left) Scanning electron micrograph of cells fixed in
the late logarithmic phase of growth; the inset shows normal E. coli cells. (Right) Thin section through E. coli cells producing Trp
polypeptide–insulin A chain fusion protein. (Photographs reproduced from Science courtesy of Dr D.C. Williams (Eli Lilly & Co.)
and the American Association for the Advancement of Science.)
Secreted proteins may be released into the periplasm
or integrated into or transported across the outer
membrane. In E. coli it has been established that protein export through the inner membrane to the
periplasm or to the outer membrane is achieved by a
universal mechanism known as the general export
pathway (GEP). This involves the sec gene products
(for review see Lory 1998). Proteins that enter the
GEP are synthesized in the cytoplasm with a signal
sequence at the N terminus. This sequence is cleaved
by a signal or leader peptidase during transport. A
signal sequence has three domains: a positively
charged amino-terminal region, a hydrophobic core,
consisting of five to 15 hydrophobic amino acids,
and a leader peptidase cleavage site. A signal sequence
attached to a normally cytoplasmic protein will
direct it to the export pathway.
Many signal sequences derived from naturally
occurring secretory proteins (e.g. OmpA, OmpT,
PelB, β-lactamase, alkaline phosphatase and phage
M13 gIII) support the efficient translocation of heterologous peptides across the inner membrane
when fused to their amino termini. In some cases,
however, the preproteins are not readily exported
and either become ‘jammed’ in the inner membrane,
accumulate in precursor inclusion bodies or are
rapidly degraded within the cytoplasm. In practice,
it may be necessary to try several signal sequences
(Berges et al. 1996) and/or overproduce different
chaperons to optimize the translocation of a particular heterologous protein. A first step would be to try
the secretion vectors offered by a number of molecularbiology suppliers and which are variants of the
vectors described above.
POGC05 9/11/2001 10:57 AM Page 84
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CHAPTER 5
It is possible to engineer proteins such that they
are transported through the outer membrane and
are secreted into the growth medium. This is achieved
by making use of the type I, Sec-independent secretion system. The prototype type I system is the
haemolysin transport system, which requires a short
carboxy-terminal secretion signal, two translocators
(HlyB and D), and the outer-membrane protein
TolC. If the protein of interest is fused to the carboxyterminal secretion signal of a type I-secreted protein,
it will be secreted into the medium provided HlyB,
HlyD and TolC are expressed as well (Spreng et al.
2000). An alternative presentation of recombinant
proteins is to express them on the surface of the
bacterial cell using any one of a number of carrier
proteins (for review, see Cornelis 2000).
Cla I 3018
T7 promoter
tac promoter
Hpa I 2777
EcoR I 3277
Pst I 3316
Sal I 52
BstX I 264
Sac I 377
purification
tag sequence
1-386
ori
PinPointTM Xa-1
Vector
(3351bp)
Factor
Xa site
379-390
Nde I
532
Nru I
Hind III
Pvu II
BamH I
Acc65 I
Kpn I
EcoR V
Bgl II
Sma I
Not I
SP6
389
392
400
404
410
414
418
422
430
435
452
Ampr
Fsp I 1483
Xmn I 1106
Pvu I 1337
Sca I 1225
Fig. 5.17 Structural features of a PinPointTM vector.
(Figure reproduced courtesy of Promega Corporation.)
Putting it all together:
vectors with combinations of features
Many of the vectors in current use, particularly
those that are commercially available, have combinations of the features described in previous sections. Two examples are described here to show the
connection between the different features. The first
example is the LITMUS vectors that were described
earlier (p. 73) and which are used for the generation
of RNA probes. They exhibit the following features:
• The polylinkers are located in the lacZα gene and
inserts in the polylinker prevent α-complementation.
Thus blue/white screening (see Box 3.2 on p. 35)
can be used to distinguish clones with inserts from
those containing vector only.
• The LITMUS polylinkers contain a total of 32
unique restriction sites. Twenty-nine of these enzymes
leave four-base overhangs and three leave blunt ends.
The three blunt cutting enzymes have been placed at
either end of the polylinker and in the middle of it.
• The vectors carry both the pUC and the M13 ori
regions. Under normal conditions the vector replicates as a double-stranded plasmid but, on infection
with helper phage (M13KO7), single-stranded molecules are produced and packaged in phage protein.
• The single-stranded molecules produced on helperphage addition have all the features necessary for
DNA sequencing (see p. 123).
• The vectors are small (< 3 kb) and with a pUC ori
have a high copy number.
The second example is the PinPoint series of
expression vectors (Fig. 5.17). These vectors have
the following features:
• Expression is under the control of either the T7
or the tac promoter, allowing the user great flexibility of control over the synthesis of the cloned gene
product.
• Some of them carry a DNA sequence specifying
synthesis of a signal peptide.
• Presence of an MCS adjacent to a factor-Xa cleavage site.
• Synthesis of an N-terminal biotinylated sequence
to facilitate purification.
• Three different vectors of each type permitting
translation of the cloned gene insert in each of the
three reading frames.
• Presence of a phage SP6 promoter distal to the
MCS to permit the synthesis of RNA probes complementary to the cloned gene. Note that the orientation of the cloned gene is known and so the RNA
probe need only be synthesized from one strand.
What is absent from these vectors is an M13 origin of
replication to facilitate synthesis of single strands for
DNA sequencing.
POGC06 9/11/2001 11:08 AM Page 85
CHAPTER 6
Cloning strategies
Introduction
In the early chapters of this book, we discussed DNA
cutting and joining techniques, and introduced
the different types of vectors that are available for
cloning DNA molecules. Any cell-based cloning
procedure has four essential parts: (i) a method
for generating the DNA fragment for cloning; (ii) a
reaction that inserts that fragment into the chosen
cloning vector; (iii) a means for introducing that
recombinant vector into a host cell wherein it is
replicated; and (iv) a method for selecting recipient
cells that have acquired the recombinant (Fig. 6.1).
To simplify the description of such procedures, the
assumption is made that we know exactly what
we are cloning. This is indeed the case with simple
subcloning strategies, where a defined restriction
fragment is isolated from one cloning vector and
inserted into another. However, we also need to
consider what happens in cases where the source
of donor DNA is very complex. We may wish, for
example, to isolate a single gene from the human
DNA
fragments
Restriction
endonuclease
digestion
Mechanical
shearing
Duplex
cDNA
synthesis
Direct
chemical
synthesis
PCR
Joining to
vector
Homopolymer
tailing
Ligation
cohesive
termini
Blunt-end
ligation
(no linker)
Linker
molecules
TA
cloning
Introduction
into host cell
Transfection
with recombinant
phage DNA
Selection or
screening
Hybridization
In vitro packaging into phage
coat: transduction with
recombinant phage or cosmid
Transformation
with recombinant
plasmid DNA
PCR
Immunochemical
Protein–protein
interactions
Protein–ligand
interactions
Functional
complementation
Gain of
function
Fig. 6.1 Generalized overview of cloning strategies, with favoured routes shown by arrows. Note that in cell-based cloning
strategies, DNA fragments are initially generated and cloned in a non-specific manner, so that screening for the desired clone is
carried out at the end of the process. Conversely, when specific DNA fragments are obtained by PCR or direct chemical synthesis,
there is no need for subsequent screening.
POGC06 9/11/2001 11:08 AM Page 86
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CHAPTER 6
genome, in which case the target sequence could be
diluted over a millionfold by unwanted genomic
DNA. We need to find some way of rapidly sifting
through, or screening, large numbers of unwanted
sequences to identify our particular target.
There are two major strategies for isolating
sequences from complex sources such as genomic
DNA. The first, a cell-based cloning strategy, is to
divide the source DNA into manageable fragments
and clone everything. Such a collection of clones,
representative of the entire starting population, is
known as a library. We must then screen the library to
identify our clone of interest, using a procedure that
discriminates between the desired clone and all the
others. A number of such procedures are discussed
later in the chapter. The second strategy is to selectively amplify the target sequence directly from the
source DNA using the polymerase chain reaction
(PCR), and then clone this individual fragment.
Each strategy has its advantages and disadvantages.
Note that, in the library approach, screening is
carried out after the entire source DNA population
has been cloned indiscriminately. Conversely, in the
PCR approach, the screening step is built into the
first stage of the procedure, when the fragments
are generated, so that only selected fragments are
actually cloned. In this chapter we consider principles for the construction and screening of genomic
and complementary DNA (cDNA) libraries, and
compare the library-based route of gene isolation to
equivalent PCR-based techniques.
Cloning genomic DNA
Genomic DNA libraries
Producing representative genomic
libraries in λ cloning vectors
Following the example above, let us suppose that we
wish to clone a single-copy gene from the human
genome. How might this be achieved? We could
simply digest total human DNA with a restriction
endonuclease, such as EcoRI, insert the fragments
into a suitable phage-λ vector and then attempt to
isolate the desired clone. How many recombinants
would we have to screen in order to isolate the right
one? Assuming EcoRI gives, on average, fragments
of about 4 kb, and given that the size of the human
haploid genome is 2.8 × 106 kb, we can see that
over 7 × 105 independent recombinants must be
prepared and screened in order to have a reasonable
chance of including the desired sequence. In other
words we have to obtain a very large number of
recombinants, which together contain a complete
collection of all of the DNA sequences in the entire
human genome, a human genomic library. The sizes
of some other genomes are listed in Table 6.1.
There are two problems with the above approach.
First, the gene may be cut internally one or more
times by EcoRI so that it is not obtained as a single
fragment. This is likely if the gene is large. Also, it
may be desirable to obtain extensive regions flanking
Table 6.1 Genome sizes of selected
organisms.
Organism
Escherichia coli
Yeast (Saccharomyces cerevisiae)
Arabidopsis thaliana (higher plant)
Tobacco
Wheat
Zea mays
Drosophila melanogaster
Mouse
Human
Xenopus laevis
Genome size (kb)
(haploid where appropriate)
4.0 × 103
1.35 × 104
1.25 × 105
1.6 × 106
5.9 × 106
1.5 × 107
1.8 × 105
2.3 × 106
2.8 × 106
3.0 × 106
POGC06 9/11/2001 11:08 AM Page 87
Cloning strategies
the gene or whole gene clusters. Fragments averaging about 4 kb are likely to be inconveniently short.
Alternatively, the gene may be contained on an
EcoRI fragment that is larger than the vector can
accept. In this case the appropriate gene would not
be cloned at all.
These problems can be overcome by cloning random DNA fragments of a large size (for λ replacement
vectors, approximately 20 kb). Since the DNA is
randomly fragmented, there will be no systematic
exclusion of any sequence. Furthermore, clones will
overlap one another, allowing the sequence of very
large genes to be assembled and giving an opportunity to ‘walk’ from one clone to an adjacent one
(p. 107). Because of the larger size of each cloned
DNA fragment, fewer clones are required for a complete or nearly complete library. How many clones
are required? Let n be the size of the genome relative
to a single cloned fragment. Thus, for the human
genome (2.8 × 106 kb) and an average cloned fragment size of 20 kb, n = 1.4 × 105. The number of
independent recombinants required in the library
must be greater than n, because sampling variation
will lead to the inclusion of some sequences several
times and the exclusion of other sequences in a
library of just n recombinants. Clarke and Carbon
(1976) have derived a formula that relates the
probability (P) of including any DNA sequence in a
random library of N independent recombinants:
N=
ln (1 − P )

1
ln 1 − 
n

Therefore, to achieve a 95% probability (P = 0.95) of
including any particular sequence in a random
human genomic DNA library of 20 kb fragment size:
N=
ln (1 − 0.95)


1
ln 1 −

5
1.4 × 10 

= 4.2 × 105
Notice that a considerably higher number of recombinants is required to achieve a 99% probability, for
here N = 6.5 × 105.
How can appropriately sized random fragments
be produced? Various methods are available. Random
breakage by mechanical shearing is appropriate
because the average fragment size can be controlled,
87
λ Charon 4A DNA
(replacement vector)
High mol. wt eukaryotic
DNA (>100 kb)
Cleave with a mixture
of HaeIII and AluI
(very partial digest)
Anneal
natural
cohesive
ends of λ
Size fractionate
~20 kb
EcoRI methylase to
block EcoRI sites
Me Me
Me Me
Blunt-end ligation with
EcoRI linker molecules
Me Me
Me Me
EcoRI
Me Me
EcoRI
+
Internal fragments
Size fractionate
to remove
internal
fragments
Me Me
Anneal EcoRI cohesive ends. Ligate
Me Me
Me Me
Me Me
Me Me
Packaging in vitro
Phage particle
Fig. 6.2 Maniatis’ strategy for producing a representative
gene library.
but insertion of the resulting fragments into vectors
requires additional steps. The more commonly used
procedure involves restriction endonucleases. In the
strategy devised by Maniatis et al. (1978) (Fig. 6.2),
the target DNA is digested with a mixture of two
restriction enzymes. These enzymes have tetranucleotide recognition sites, which therefore occur
frequently in the target DNA and in a complete
double-digest would produce fragments averaging
less than 1 kb. However, only a partial restriction
digest is carried out, and therefore the majority of
the fragments are large (in the range 10–30 kb).
Given that the chances of cutting at each of the
available restriction sites are more or less equivalent, such a reaction effectively produces a
random set of overlapping fragments. These can be
size-fractionated, e.g. by gel electrophoresis, so as
to give a random population of fragments of about
POGC06 9/11/2001 11:08 AM Page 88
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CHAPTER 6
Polylinker
BamHI
SalI
EcoRI
red+
AB
SalI
Polylinker
BamHI
EcoRI SalI
gam+
λEMBL3A
Target high molecular
weight genomic DNA
SalI
(1) Partial Sau3AI digest
(2) Phosphatase
Digest with BamHI
and EcoRI
AB
BamHI red+
gam+
EcoRI
Left arm
Polylinker
oligonucleotide
EcoRI
BamHI
BamHI
HO
Sau3AI
Sau3AI
OH HO
Sau3AI
BamHI
EcoRI
Left arm
Sau3AI Sau3AI Sau3AI
OH HO
OH
Sau3AI Sau3AI
Isopropanol precipitation
Polylinker oligonucleotide
not precipitated
AB
Sau3AI
HO
Right arm
Polylinker
oligonucleotide
OH
Sau3AI
HO
OH
Size fractionation not necessary
Right arm
Ligate, mix
AB
(BamHI)
(BamHI)
Left arm
Right arm
SalI
+
Recombinant phage
DNA concatemers
SalI
Packaging in vitro
Recombinant
(Spi–) phage
plaques
Plate on
P2 lysogen
sup+
Fig. 6.3 Creation of a genomic DNA library using the phage-λ vector EMBL3A. High-molecular-weight genomic DNA is partially
digested with Sau3AI. The fragments are treated with phosphatase to remove their 5′ phosphate groups. The vector is digested
with BamHI and EcoRI, which cut within the polylinker sites. The tiny BamHI/EcoRI polylinker fragments are discarded in the
isopropanol precipitation, or alternatively the vector arms may be purified by preparative agarose gel electrophoresis. The vector
arms are then ligated with the partially digested genomic DNA. The phosphatase treatment prevents the genomic DNA fragments
from ligating together. Non-recombinant vector cannot re-form because the small polylinker fragments have been discarded.
The only packageable molecules are recombinant phages. These are obtained as plaques on a P2 lysogen of sup+ E. coli. The Spi−
selection ensures recovery of phage lacking red and gam genes. A sup+ host is necessary because, in this example, the vector carries
amber mutations in genes A and B. These mutations increase biological containment, and can be applied to selection procedures,
such as recombinational selection, or tagging DNA with a sup+ gene. Ultimately, the foreign DNA can be excised from the vector
by virtue of the SalI sites in the polylinker. (Note: Rogers et al. (1988) have shown that the EMBL3 polylinker sequence is not
exactly as originally described. It contains an extra sequence with a previously unreported PstI site. This does not affect most
applications as a vector.)
POGC06 9/11/2001 11:08 AM Page 89
Cloning strategies
20 kb, which are suitable for insertion into a λ replacement vector. Packaging in vitro (p. 58) ensures
that an appropriately large number of independent
recombinants can be recovered, which will give an
almost completely representative library.
Development of λ replacement vectors for
genomic library construction
In the Maniatis strategy, the use of two different
restriction endonucleases with completely unrelated
recognition sites, HaeIII and AluI, assists in obtaining
fragmentation that is nearly random. These enzymes
both produce blunt ends, and the cloning strategy
requires linkers (see Fig. 6.2). Therefore, in the early
days of vector development, a large number of
different vectors became available with alternative
restriction sites and genetic markers suitable for
varied cloning strategies. A good example of this
diversity is the Charon series, which included both
insertion- and replacement-type vectors (Blatttner
et al. 1977, Williams & Blattner 1979).
A convenient simplification can be achieved by
using a single restriction endonuclease that cuts frequently, such as Sau3AI. This will create a partial
digest that is slightly less random than that achieved
with a pair of enzymes. However, it has the great
advantage that the Sau3AI fragments can be readily
inserted into λ replacement vectors, such as λEMBL3
(Frischauf et al. 1983), which have been digested
with BamHI (Fig. 6.3). This is because Sau3AI and
BamHI create the same cohesive ends (see p. 32). Due
to the convenience and efficiency of this strategy,
the λEMBL series of vectors have been very widely
used for genomic library construction (p. 58). Note
that λEMBL vectors also carry the red and gam genes
on the stuffer fragment and a chi site on one of the
vector arms, allowing convenient positive selection on the basis of the Spi phenotype (see p. 58).
Most λ vectors currently used for genomic library
construction are positively selected on this basis,
including λ2001 (Karn et al. 1984), λDASH and
λFIX (Sorge 1988). λDASH and λFIX (and recently
improved versions, λDASHII and λFIXII) are particularly versatile because the multiple cloning sites
flanking the stuffer fragment contain opposed
promoters for the T3 and T7 RNA polymerases. If
the recombinant vector is digested with a restriction
89
endonuclease that cuts frequently, only short fragments of insert DNA are left attached to these
promoters. This allows RNA probes to be generated
corresponding to the ends of any genomic insert.
These are ideal for probing the library to identify
overlapping clones as part of a chromosome walk
(p. 107) and have the great advantage that they can
be made conveniently, directly from the vector, without recourse to subcloning. Vector maps of λDASH
and λFIX are shown in Fig. 6.4. λFIX is similar to
λDASH, except that it incorporates additional XhoI
sites flanking the stuffer fragment. Digestion of the
vector with XhoI followed by partial filling of the
sticky ends prevents vector religation. However,
partially filled Sau3AI sticky ends are compatible
with the partially filled XhoI ends, although not with
each other. This strategy prevents the ligation of
vector arms without genomic DNA, and also prevents
the insertion of multiple fragments.
Genomic libraries in high-capacity vectors
In place of phage-λ derivatives, a number of highercapacity cloning vectors such as cosmids, bacterial
artificial chromosomes (BACs), P1-derived artificial
chromosomes (PACs) and yeast artificial chromosomes (YACs) are available for the construction of
genomic libraries. The advantage of such vectors
is that the average insert size is much larger than
for λ replacement vectors. Thus, the number of
recombinants that need to be screened to identify a
particular gene of interest is correspondingly lower,
large genes are more likely to be contained within
a single clone and fewer steps are needed for a chromosome walk (p. 107). Generally, strategies similar
to the Maniatis method discussed above are used
to construct such libraries, except that the partial
restriction digest conditions are optimized for larger
fragment sizes, and size fractionation must be preformed by specialized electrophoresis methods that
can separate fragments over 30 kb in length. Pulsedfield gel electrophoresis (PFGE) and field-inversion
gel electrophoresis (FIGE) have been devised for this
purpose (p. 10). High-molecular-weight donor DNA
fragments can also be prepared using restriction
enzymes that cut very rarely.
Cosmids may be favoured over λ vectors because
they accept inserts of up to 45 kb. However, since
POGC06 9/11/2001 11:08 AM Page 90
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CHAPTER 6
SalI
Xba I
N
C II
gam+
gam+
N
14 kb
(b)
these vectors are maintained as high-copy-number
plasmids in Escherichia coli, they have a tendency
to be unstable, undergoing deletions that favour
increased replication. Most workers also find that
plaques give less background hybridization than
colonies, so screening libraries of phage-λ recombinants by plaque hybridization gives cleaner results
than screening cosmid libraries by colony hybridization (see p. 104). It may also be desired to retain and
store an amplified genomic library. With phage, the
initial recombinant DNA population is packaged
and plated out, and can be screened at this stage.
Alternatively, the plates containing the recombinant plaques can be washed to give an amplified library
of recombinant phage. The amplified library can
then be stored almost indefinitely, because phage
have a long shelf-life. The amplification is so great
that samples of the amplified library can be plated
out and screened with different probes on hundreds
of occasions. With bacterial colonies containing
cosmids, it is also possible to store an amplified
library, but bacterial populations cannot be stored
as readily as phage populations, and there is often
an unacceptable loss of viability (Hanahan &
Meselson 1980). A word of caution is necessary,
however, when considering the use of any amplified
library. This is due to the possibility of distortion.
Not all recombinants in a population will propagate
C II
red+
SacI
Xba I
Xba I
EcoRI
XhoI
SalI
Xba I
SacI
J
20 kb
λDASH
T3
SalI
XhoI
EcoRI
Xba I
T7
A
O P QS R
9 kb
14 kb
cro
20 kb
cro
red+
J
(a)
Xba I
XhoI
SacI
HindIII
BamHI
EcoRI
Xba I
SalI
A
T7
EcoRI
BamHI
HindIII
SacI
XhoI
Xba I
T3
O P QS R
9 kb
λFIX
Fig. 6.4 The replacement vectors
λDASH and λFIX. Promoters
specific for the bacteriophage T3
and T7 RNA polymerases are
located adjacent to the cloning
sites, allowing RNA probes to be
generated that correspond to
each end of the insert.
equally well, e.g. variations in target DNA size or
sequence may affect replication of a recombinant
phage, plasmid or cosmid. Therefore, when a library
is put through an amplification step, particular
recombinants may be increased in frequency,
decreased in frequency or lost altogether. The development of modern vectors and cloning strategies
has simplified library construction to the point
where many workers now prefer to create a new
library for each screening, rather than risk using a
previously amplified one. Furthermore, pre-made
libraries are available from many commercial
sources and the same companies often offer custom
library services. These libraries are often of high
quality and such services are becoming increasingly
popular.
The highest-capacity vectors – BACs, PACs and
YACs – would seem to be ideal for library construction because of the very large insert sizes.
However, such libraries are generally more difficult
to prepare, and the larger inserts can be less than
straightforward to work with. Unless the genomic
target sequence is known to be very large and needs
to be isolated as a single clone, λ replacement vectors
or cosmids remain the most appropriate choice for
many experiments. The main application of BAC,
PAC and YAC libraries is for genome mapping,
sequencing and the assembly of clone contigs.
POGC06 9/11/2001 11:08 AM Page 91
Cloning strategies
Subgenomic libraries
Genomic libraries have been prepared from single
human chromosomes separated by flow cytometry
(e.g. Davies et al. 1981). Even greater enrichment for
particular regions of the genome is possible using
the technically demanding technique of chromosome
microdissection. In Drosophila, it has been possible
to physically excise a region of the salivary gland
chromosome by micromanipulation, and then digest
the DNA and clone it in phage-λ vectors, all within
a microdrop under oil (Scalenghe et al. 1981).
Similarly, this has been achieved with specific bands
of human chromosomes using either extremely fine
needles or a finely focused laser beam (Ludecke et al.
1989, 1990). Regardless of the species, this is a
laborious and difficult technique and is prone to
contamination with inappropriate DNA fragments.
It has been rendered obsolete with the advent of
high-capacity vectors, such as YACs.
PCR as an alternative to genomic
DNA cloning
The PCR is a robust technique for amplifying specific
DNA sequences from complex sources. In principle,
therefore, PCR with specific primers could be used to
isolate genes directly from genomic DNA, obviating
the need for the production of genomic libraries.
However, a serious limitation is that standard PCR
conditions are suitable only for the amplification of
short products. The maximum product size that
can be obtained is about 5 kb, although the typical
size is more likely to be 1–2 kb. This reflects the
poor processivity of PCR enzymes such as Taq polymerase, and their lack of proofreading activity. Both
of these deficiencies increase the likelihood of the
enzyme detaching from the template, especially if
the template is long. The extreme reaction conditions required for the PCR are also thought to
cause damage to bases and generate nicks in DNA
strands, which increases the probability of premature termination on long templates.
Long PCR
Modifications to reaction conditions can improve
polymerase processivity by lowering the reaction
91
temperature and increasing the pH, thereby protecting the template from damage (Foord & Rose 1994).
The use of such conditions in combination with two
DNA polymerases, one of which is a proofreading
enzyme, has been shown to dramatically improve
the performance of PCR using long templates (Barnes
1994, Cheng et al. 1994a). Essentially, the improvements come about because the proofreading enzyme
removes mismatched bases that are often incorporated into growing DNA strands by enzymes such
as Taq polymerase. Under normal conditions, Taq
polymerase would stall at these obstructions and,
lacking the intrinsic proofreading activity to correct
them, the reaction would most likely be aborted.
Using such polymerase mixtures, it has been
possible to amplify DNA fragments of up to 22 kb
directly from human genomic DNA, almost the
entire 16.6 kb human mitochondrial genome and
the complete or near-complete genomes of several
viruses, including 42 kb of the 45 kb phage-λ
genome (Cheng et al. 1994a,b). Several commercial
companies now provide cocktails of enzymes suitable for long PCR, e.g. TaqPlus Long PCR system,
marketed by Stratagene, which is essentially a
mixture of Taq polymerase and the thermostable
proofreading enzyme Pfu polymerase. The technique has been applied to the structural analysis
of human genes (e.g. Ruzzo et al. 1998, Bochmann
et al. 1999) and viral genomes, including human
immunodeficiency virus (HIV) (Dittmar et al. 1997).
Long PCR has particular diagnostic value for the
analysis of human triplet-repeat disorders, such as
Friedreich’s ataxia (Lamont et al. 1997). However,
while long PCR is useful for the isolation of genes
where sequence information is already available, it
is unlikely to replace the use of genomic libraries,
since the latter represent a permanent, full-genome
resource that can be shared by numerous laboratories. Indeed, genomic libraries may be used in preference to total genomic DNA as the starting-point for
gene isolation by long PCR.
Fragment libraries
Traditional genomic libraries cannot be prepared
from small amounts of starting material, e.g. single
cells, or from problematical sources, such as fixed tissue. In these cases, PCR is the only available strategy
POGC06 9/11/2001 11:08 AM Page 92
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CHAPTER 6
for gene isolation. However, as well as being useful
for the isolation of specific fragments, the PCR can be
used to generate libraries, i.e. by amplifying a representative collection of random genomic fragments.
This can be achieved using either random primers
followed by size selection for suitable PCR products
or a strategy in which genomic DNA is digested with
restriction enzymes and then linkers are ligated to
the ends of the DNA fragments, providing annealing
sites for one specific type of primer (e.g. Zhang et al.
1992, Cheung & Nelson 1996). These techniques
are powerful because they allow genomic fragment
libraries to be prepared from material that could not
yield DNA of suitable quality or quantity for conventional library construction, but competition among
the templates generally does not allow the production of a truly representative library.
cDNA cloning
Properties of cDNA
cDNA is prepared by reverse-transcribing cellular
RNA. Cloned eukaryotic cDNAs have their own
special uses, which derive from the fact that they
lack introns and other non-coding sequences present in the corresponding genomic DNA. Introns
are rare in bacteria but occur in most genes of
higher eukaryotes. They can be situated within the
coding sequence itself, where they then interrupt
the collinear relationship of the gene and its encoded
polypeptide, or they may occur in the 5′ or 3′ untranslated regions. In any event, they are copied by RNA
polymerase when the gene is transcribed. The
primary transcript goes through a series of processing events in the nucleus before appearing in the
cytoplasm as mature mRNA. These events include
the removal of intron sequences by a process called
splicing. In mammals, some genes contain numerous large introns that represent the vast majority of
the sequence. For example, the human dystrophin
gene contains 79 introns, representing over 99% of
the sequence. The gene is nearly 2.5 Mb in length
and yet the corresponding cDNA is only just over 11
kb. Thus, one advantage of cDNA cloning is that in
many cases the size of the cDNA clone is significantly
lower than that of the corresponding genomic clone.
Since removal of eukaryotic intron transcripts by
splicing does not occur in bacteria, eukaryotic cDNA
clones find application where bacterial expression of
the foreign DNA is necessary, either as a prerequisite
for detecting the clone (see p. 109) or because expression of the polypeptide product is the primary
objective. Also, where the sequence of the genomic
DNA is available, the position of intron/exon boundaries can be assigned by comparison with the cDNA
sequence.
cDNA libraries
Under some circumstances, it may be possible to prepare cDNA directly from a purified mRNA species.
Much more commonly, a cDNA library is prepared
by reverse-transcribing a population of mRNAs and
then screened for particular clones. An important
concept is that the cDNA library is representative of
the RNA population from which it was derived.
Thus, whereas genomic libraries are essentially the
same, regardless of the cell type or developmental
stage from which the DNA was isolated, the contents of cDNA libraries will vary widely according
to these parameters. A given cDNA library will also
be enriched for abundant mRNAs but may contain
only a few clones representing rare mRNAs. Furthermore, where a gene is differentially spliced, a
cDNA library will contain different clones representing alternative splice variants.
Table 6.2 shows the abundances of different classes
of mRNAs in two representative tissues. Generally,
mRNAs can be described as abundant, moderately
abundant or rare. Notice that, in the chicken
oviduct, one mRNA type is superabundant. This
encodes ovalbumin, the major egg-white protein.
Therefore, the starting population is naturally so
enriched in ovalbumin mRNA that isolating the
ovalbumin cDNA can be achieved without the use of
a library. An appropriate strategy for obtaining such
abundant cDNAs is to clone them directly in an M13
vector, such as M13 mp8. A set of clones can then be
sequenced immediately and identified on the basis
of the polypeptide that each encodes. A successful
demonstration of this strategy was reported by
Putney et al. (1983), who determined DNA sequences
of 178 randomly chosen muscle cDNA clones. Based
on the amino acid sequences available for 19
abundant muscle-specific proteins, they were able to
POGC06 9/11/2001 11:08 AM Page 93
Cloning strategies
93
Table 6.2 Abundance classes of typical
mRNA populations.
Number of
different mRNAs
Source
Abundance
(molecules/cell)
Mouse liver cytoplasmic poly(A)+
1
2
3
9
700
11 500
12 000
300
15
Chick oviduct polysomal poly(A)+
1
2
3
1
7
12 500
100 000
4 000
5
References: mouse (Young et al. 1976); chick oviduct (Axel et al. 1976).
identify clones corresponding to 13 of these 19
proteins, including several protein variants.
For the isolation of cDNA clones in the moderateand low-abundance classes, it is usually necessary
to construct a cDNA library. Once again, the high
efficiency obtained by packaging in vitro makes
phage-λ vectors attractive for obtaining large numbers of cDNA clones. Phage-λ insertion vectors are
particularly well suited for cDNA cloning and some of
the most widely used vectors are discussed in Box 6.1.
Typically, 105–106 clones are sufficient for the isolation of low-abundance mRNAs from most cell types,
i.e. those present at 15 molecules per cell or above.
However, some mRNAs are even less abundant than
this, and may be further diluted if they are expressed
in only a few specific cells in a particular tissue. Under
these circumstances, it may be worth enriching the
mRNA preparation prior to library construction, e.g.
by size fractionation, and testing the fractions for the
presence of the desired molecule. One way in which
Box 6.1 Phage-l vectors for cDNA cloning and expression
lgt10 and lgt11
Most early cDNA libraries were constructed using
plasmid vectors, and were difficult to store and
maintain for long periods. They were largely replaced
by phage-l libraries, which can be stored indefinitely
and can also be prepared to much higher titres.
lgt10 and lgt11 were the standard vectors for
cDNA cloning until about 1990. Both lgt10 and
lgt11 are insertion vectors, and they can accept
approximately 7.6 kb and 7.2 kb of foreign DNA,
respectively. In each case, the foreign DNA is
introduced at a unique EcoRI cloning site. lgt10
is used to make libraries that are screened by
hybridization. The EcoRI site interrupts the phage
cI gene, allowing selection on the basis of plaque
morphology. lgt11 contains an E. coli lacZ gene
driven by the lac promoter. If inserted in the correct
orientation and reading frame, cDNA sequences
cloned in this vector can be expressed as
b-galactosidase fusion proteins, and can be detected
by immunological screening or screening with other
ligands (see p. 109). lgt11 libraries can also be
screened by hybridization, although lgt10 is more
appropriate for this screening strategy because
higher titres are possible.
lZAP series
While phage-l vectors generate better libraries, they
cannot be manipulated in vitro with the convenience
of plasmid vectors. Therefore, phage clones have to
continued
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CHAPTER 6
Box 6.1 continued
be laboriously subcloned back into plasmids for
further analysis. This limitation of conventional
phage-l vectors has been addressed by the
development of hybrids, sometimes called phasmids,
which possess the most attractive features of both
bacteriophage l and plasmids (see Chapter 5). The
most popular current vectors for cDNA cloning are
undoubtedly those of the lZAP series marketed by
Stratagene (Short et al. 1988). A map of the original
lZAP vector is shown below. The advantageous
features of this vector are: (i) the high capacity – up
to 10 kb of foreign DNA can be cloned, which is large
enough to encompass most cDNAs; (ii) the presence
of a polylinker with six unique restriction sites,
which increases cloning versatility and also allows
directional cloning; and (iii) the availability of T3
and T7 RNA polymerase sites flanking the polylinker,
allowing sense and antisense RNA to be prepared
from the insert. Most importantly, all these features
are included within a plasmid vector called
pBluescript, which is itself inserted into the phage
genome. Thus the cDNA clone can be recovered from
the phage and propagated as a high-copy-number
plasmid without any subcloning, simply by
coinfecting the bacteria with a helper f1 phage
that nicks the lZAP vector at the flanks of the
plasmid and facilitates excision. Another member
of this series, lZAP Express, also includes the human
cytomegalovirus promoter and SV40 terminator,
so that fusion proteins can be expressed in
mammalian cells as well as bacteria. Thus, cDNA
libraries can be cloned in the phage vector in E. coli,
rescued as plasmids and then transfected into
mammalian cells for expression cloning.
T3
I
T7
p Bluescript
T
cos
cos
f1 origin
T
Ampr
I
hc2
ColE1ori
T7
T3
POGC06 9/11/2001 11:08 AM Page 95
Cloning strategies
this can be achieved is to inject mRNA fractions into
Xenopus oocytes (p. 215) and test them for production of the corresponding protein (Melton 1987). See
also the discussion of subtraction cloning on p. 115.
Preparation of cDNA for library
construction
The cDNA synthesis reaction
The synthesis of double-stranded cDNA suitable for
insertion into a cloning vector involves three major
steps: (i) first-strand DNA synthesis on the mRNA
template, carried out with a reverse transcriptase;
(ii) removal of the RNA template; and (iii) secondstrand DNA synthesis using the first DNA strand as a
template, carried out with a DNA-dependent DNA
polymerase, such as E. coli DNA polymerase I. All
DNA polymerases, whether they use RNA or DNA
as the template, require a primer to initiate strand
synthesis.
Development of cDNA cloning strategies
The first reports of cDNA cloning were published in
the mid-1970s and were all based on the homopolymer tailing technique, which is described briefly in
Chapter 3. Of several alternative methods, the one
that became the most popular was that of Maniatis
et al. (1976). This involved the use of an oligo-dT
primer annealing at the polyadenylate tail of the
mRNA to prime first-strand cDNA synthesis, and
took advantage of the fact that the first cDNA strand
has the tendency to transiently fold back on itself,
forming a hairpin loop, resulting in self-priming of
the second strand (Efstratiadis et al. 1976). After the
synthesis of the second DNA strand, this loop must
be cleaved with a single-strand-specific nuclease,
e.g. S1 nuclease, to allow insertion into the cloning
vector (Fig. 6.5).
A serious disadvantage of the hairpin method is
that cleavage with S1 nuclease results in the loss of
a certain amount of sequence at the 5′ end of the
clone. This strategy has therefore been superseded
by other methods in which the second strand is
primed in a separate reaction. One of the simplest
strategies is shown in Fig. 6.6 (Land et al. 1981).
After first-strand synthesis, which is primed with an
95
oligo-dT primer as usual, the cDNA is tailed with a
string of cytidine residues using the enzyme terminal transferase. This artificial oligo-dC tail is then
used as an annealing site for a synthetic oligo-dG
primer, allowing synthesis of the second strand.
Using this method, Land et al. (1981) were able to
isolate a full-length cDNA corresponding to the
chicken lysozyme gene. However, the efficiency can
be lower for other cDNAs (e.g. Cooke et al. 1980).
For cDNA expression libraries, it is advantageous
if the cDNA can be inserted into the vector in the
correct orientation. With the self-priming method,
this can be achieved by adding a synthetic linker
to the double-stranded cDNA molecule before the
hairpin loop is cleaved (e.g. Kurtz & Nicodemus
1981; Fig. 6.7a). Where the second strand is primed
separately, direction cloning can be achieved using
an oligo-dT primer containing a linker sequence
(e.g. Coleclough & Erlitz 1985; Fig. 6.7b). An alternative is to use primers for cDNA synthesis that are
already linked to a plasmid (Fig. 6.7c). This strategy
was devised by Okayama and Berg (1982) and has
two further notable characteristics. First, full-length
cDNAs are preferentially obtained because an RNA–
DNA hybrid molecule, the result of first-strand
synthesis, is the substrate for a terminal transferase
reaction. A cDNA that does not extend to the end of
the mRNA will present a shielded 3-hydroxyl group,
which is a poor substrate for tailing. Secondly, the
second-strand synthesis step is primed by nicking the
RNA at multiple sites with RNase H. Second-strand
synthesis therefore occurs by a nick-translation
type of reaction, which is highly efficient. Simpler
cDNA cloning strategies incorporating replacement
synthesis of the second strand are widely used (e.g.
Gubler & Hoffman 1983, Lapeyre & Amalric 1985).
The Gubler–Hoffman reaction, as it is commonly
known, is show in Fig. 6.8.
Full-length cDNA cloning
Limitations of conventional cloning strategies
Conventional approaches to the production of cDNA
libraries have two major drawbacks. First, where
oligo-dT primers are used to initiate first-strand synthesis, there is generally a 3′-end bias (preferential
recovery of clones representing the 3′ end of cDNA
POGC06 9/11/2001 11:08 AM Page 96
96
CHAPTER 6
mRNA
5’
AAAn 3’
Oligo (dT)
AAAn 3’
TTTn
5’
TcR
PstI
(1) Reverse
transcriptase
(2) Alkali
pBR322
ApR
First strand synthesis
TTTn 5’
cDNA
3’
PstI
Second strand synthesis
G
3’ ACGTC
DNA polymerase
+ 4 dNTPs
5’
3’
S1-nuclease
CTGCA 3’
G
3’
Terminal
transferase
+ dGTP
G
GGnGACGTC
Terminal
transferase
+ dCTP
CTGCAGGnG
G
3’
CCnC
CCnC
Annealing
cDNA
G
CCnC
CTGCAGGnG
GGnGACGTC
CCnG
G
Select
TcR, ApS
GACGTCCnC
CTGCAGGnG
Transformation host repairs
gaps, reconstructs PstI sites
GGnGACGTC
CCnCTGCAG
Fig. 6.5 An early cDNA cloning strategy, involving hairpin-primed second-strand DNA synthesis and homopolymer tailing to
insert the cDNA into the vector.
sequences) in the resulting library. This can be
addressed through the use of random oligonucleotide primers, usually hexamers, for both first- and
second-strand cDNA synthesis. However, while this
eliminates 3′-end bias in library construction, the
resulting clones are much smaller, such that full-
length cDNAs must be assembled from several
shorter fragments. Secondly, as the size of a cDNA
increases, it becomes progressively more difficult to
isolate full-length clones. This is partly due to
deficiencies in the reverse-transcriptase enzymes
used for first-strand cDNA synthesis. The enzymes
POGC06 9/11/2001 11:08 AM Page 97
Cloning strategies
97
mRNA
5‘
A A A A 3‘
Oligo-dT
Reverse transcriptase
+ 4 dNTPs
First strand synthesis
mRNA
5‘
3‘
A A A A 3‘
T T T T 5‘
cDNA
Terminal transferase
+ dCTP
mRNA
5‘
3‘ C C C C
cDNA
A A A A CCC
3‘
TTT
5‘
Alkaline sucrose gradient:
(1) hydrolyses RNA
(2) recovers full length cDNA
3‘ C C C C
cDNA
T T T T 5‘
Oligo-dG, reverse transcriptase
+ 4 dNTPs
Second strand synthesis
5‘
GGG
3‘
3‘ C C C C
T T T T 5‘
Fig. 6.6 Improved method for cDNA
cloning. The first strand is tailed with
oligo(dC) allowing the second strand to
be initiated using an oligo(dG) primer.
are usually purified from avian myeloblastosis virus
(AMV) or produced from a cloned Moloney murine
leukaemia virus (MMLV) gene in E. coli. Native
enzymes have poor processivity and intrinsic RNase
activity, which leads to degradation of the RNA template (Champoux 1995). Several companies produce engineered murine reverse transcriptases that
lack RNase H activity, and these are more efficient
in the production of full-length cDNAs (Gerard &
D’Allesio 1993). An example is the enzyme SuperScript II, marketed by Life Technologies (Kotewicz
et al. 1988). This enzyme can also carry out reverse
transcription at temperatures of up to 50°C. The native
enzymes function optimally at 37°C and therefore
tend to stall at sequences that are rich in secondary
duplex cDNA
Insert into vector by
either further homopolymer
tailing or linkers
structure, as often found in 5′ and 3′ untranslated
regions.
Selection of 5′ mRNA ends
Despite improvements in reverse transcriptases,
the generation of full-length clones corresponding
to large mRNAs remains a problem. This has been
addressed by the development of cDNA cloning
strategies involving the selection of mRNAs with
intact 5′ ends. Nearly all eukaryotic mRNAs have
a 5′ end cap, a specialized, methylated guanidine
residue that is inverted with respect to the rest of the
strand and is recognized by the ribosome prior to the
initiation of protein synthesis. Using a combination
POGC06 9/11/2001 11:08 AM Page 98
98
CHAPTER 6
(a)
(b)
mRNA
5‘
A A An 3‘
mRNA
5‘
Oligo-dT
A A An
T T Tn
5‘
5‘
S
(1) Reverse transcriptase + 4dNTPs
(1) Reverse transcriptase + 4dNTPs
(2) Alkali
First strand synthesis
First strand synthesis
T T Tn 5‘
cDNA
A A An 3‘
Oligo-dT (SalI primer)
A A An
T T Tn
T T Tn
3‘
3‘
S
DNA polymerase (Klenow) + 4dNTPs
DNA polymerase (Klenow) + 4dNTPs
Second strand synthesis
Second strand synthesis
A A An
SalI-linkers, ligation
S S S
A A An
E
Cut with SalI, EcoRI
(1) Nuclease S1
(2) DNA polymerase (Klenow)
+ 4dNTPs
S
T T Tn
EcoRI linkers
E
S
E
T T Tn
S
S S S
Insert into vector cut with SalI and EcoRI
EcoRI-linkers, ligation
R R
S S S R R
Cut with SalI, EcoRI
R
S
Insert into vector cut with SalI and EcoRI
(c)
mRNA
– – – – – – –AAAAAA
HindIII
AmpR
Primer
T
T
T
T
CC
Annealing
cDNA
synthesis
PvuII First
strand
synthesis
A
T AA
T
T A
T
Annealing of oligo-dG
tailed adaptor;
cyclization with HindIII
E. coli DNA ligase
G
G
PstI G CC
C
HindIII
A
T A
T
TA
TA
PvuII
Second
strand
synthesis
PvuII
HindIII
PstI
GG
G
Replacement of
RNA strand by
DNA with E. coli
RNase H, DNA
polymerase I and
DNA ligase
C
OligodC tails
CC
C
A
T AA
T
T A
T
HindIII
digestion
HindIII
CCC
G
G C
G C
C
HindIII
A
T A
T
TA
TA
HindIIIC
C
HindIII
C
A
T AA
T
T A
T
PvuII
POGC06 9/11/2001 11:08 AM Page 99
Cloning strategies
99
A A A An mRNA
cap
Oligo (dT)
A A A An mRNA
T T T Tn cDNA
Reverse transcriptase
+ 4 dNTPs
A A A An mRNA
T T T Tn cDNA
RNAseH
random primers
DNA polymerase
AAA
First strand
cDNA synthesis
Full
length
cDNA
cap
RNAse A
treatment
T T T Tn cDNA
A A A An duplex
T T T Tn cDNA
Fig. 6.8 The Gubler–Hoffman method, a simple and general
method for non-directional cDNA cloning. First-strand
synthesis is primed using an oligo(dT) primer. When the first
strand is complete, the RNA is removed with RNase H and the
second strand is random-primed and synthesized with DNA
polymerase I. T4 DNA polymerase is used to ensure that the
molecule is blunt-ended prior to insertion into the vector.
of cap selection and nuclease treatment, it is possible
to select for full-length first-strand cDNAs and thus
generate libraries highly enriched in full-length
clones.
An example is the method described by Edery et al.
(1995) (Fig. 6.9). In this strategy, first-strand cDNA
synthesis is initiated as usual, using an oligo-dT
primer. Following the synthesis reaction, the hybrid
molecules are treated with RNase A, which only
Fig. 6.7 (opposite) Methods for directional cDNA cloning. (a)
An early strategy in which the formation of a loop is exploited
to place a specific linker (in this example, for SalI) at the open
end of the duplex cDNA. Following this ligation, the loop is
cleaved and trimmed with S1 nuclease and EcoR1 linkers are
added to both ends. Cleavage with EcoR1 and SalI generates
a restriction fragment that can be unidirectionally inserted
into a vector cleaved with the same enzymes. (b) A similar
strategy, but second-strand cDNA synthesis is randomprimed. The oligo(dT) primer carries an extension forming
a SalI site. During second-strand synthesis, this forms a
double-stranded SalI linker. The addition of further EcoRI
linkers to both ends allows the cDNA to be unidirectionally
cloned, as above. (c) The strategy of Okayama and Berg
(1982), where the mRNA is linked unidirectionally to the
plasmid cloning vector prior to cDNA synthesis, by virtue
of a cDNA tail.
Partial
length
cDNA
AAA
TTT
cap
cap
AAA
TTT
RNAse A
treatment
AAA
TTT
Isolation on
elF-4E affinity
column
AAA
TTT
melf-4E
A
cap
AAA
TTT
Elute full length cDNA
Fig. 6.9 The CAPture method of full-length cDNA cloning,
using the eukaryotic initiation factor eIF-4E to select
mRNAs with caps protected from RNase digestion by a
complementary DNA strand.
digests single-stranded RNA. DNA–RNA hybrids
therefore remain intact. If the first-strand cDNA is
full-length, it reaches all the way to the 5′ cap of the
mRNA, which is therefore protected from cleavage
by RNase A. However, part-length cDNAs will leave
a stretch of unprotected single-stranded RNA between
the end of the double-stranded region and the cap,
which is digested away with the enzyme. In the next
stage of the procedure, the eukaryotic translational
initiation factor eIF-4E is used to isolate full-length
molecules by affinity capture. Incomplete cDNAs
and cDNAs synthesized on broken templates will
lack the cap and will not be retained. A similar
method based on the biotinylation of mRNA has also
been reported (Caminci et al. 1996). Both methods,
however, also co-purify cDNAs resulting from the
mispriming of first-strand synthesis, which can
account for up to 10% of the clones in a library.
An alternative method, oligo-capping, addresses this
POGC06 9/11/2001 11:08 AM Page 100
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CHAPTER 6
Oligo-capping
Full-length mRNA cap
Partial-length mRNA
AAAAAAA
P
AAAAAAA
BAP treatment
cap
AAAAAAA
OH
AAAAAAA
TAP treatment
P
AAAAAAA
OH
AAAAAAA
Oligo plus RNA
ligase
Oligo capped
mRNA
AAAAAAA
OH
AAAAAAA
First strand
synthesis
AAAAAAA
TTTTTTT
OH
AAAAAAA
TTTTTTT
PCR amplification and cloning
(only molecules containing both
primers will be amplified)
AAAAAAA
TTTTTTT
Fig. 6.10 Oligo-capping, the addition of specific
oligonucleotide primers to full-length RNAs by
sequential treatment with alkaline phosphatase and
acid pyrophosphatase. Once the oligo cap has annealed
to the 5′ end of the mRNA, it can serve as a primer binding
site for PCR amplification.
problem by performing selection at the RNA stage
(Maruyama & Sugano 1994, Suzuki et al. 1997,
2000; Fig. 6.10). The basis of the method is that
RNA is sequentially treated with the enzymes
alkaline phosphatase and acid pyrophosphatase.
The first enzyme removes phosphate groups from
the 5′ ends of uncapped RNA molecules, but does
not affect full-length molecules with a 5′ cap. The
second treatment removes the cap from full-length
RNAs, leaving a 5′-terminal residue with a phosphate
group. Full-length molecules can be ligated to a
specific oligonucleotide, while broken and degraded
molecules cannot. The result is an oligo-capped
population of full-length mRNAs. This selected population is then reverse-transcribed using an oligo-dT
primer. Second-strand synthesis and cloning is then
carried out by PCR using the oligo-dT primer and
a primer annealing to the oligonucleotide cap. Only
full-length cDNAs annealing to both primers will
be amplified, thus eliminating broken or degraded
RNAs, incomplete first cDNA strands (which lack
a 5′ primer annealing site) and misprimed cDNAs
(which lack a 3′ primer annealing site).
PCR as an alternative to cDNA cloning
Reverse transcription followed by the PCR (RT-PCR)
leads to the amplification of RNA sequences in cDNA
form. No modification to the basic PCR strategy
(p. 19) is required, except that the template for
PCR amplification is generated in the same reaction
tube in a prior reverse-transcription reaction (see
Kawasaki 1990, Dieffenbach & Dvesler 1995). Using
gene-specific primers, RT-PCR is a sensitive means
for detecting, quantifying and cloning specific cDNA
molecules. Reverse transcription is carried out using
a specific 3′ primer that generates the first cDNA
strand, and then PCR amplification is initiated following the addition of a 5′ primer to the reaction mix.
The sensitivity is such that total RNA can be used
as the starting material, rather than the poly(A)+
RNA which is used for conventional cDNA cloning.
Total RNA also contains ribosomal RNA (rRNA)
and transfer RNA (tRNA), which can be present in a
great excess to mRNA.
Due to the speed with which RT-PCR can be
carried out, it is an attractive approach for obtaining
a specific cDNA sequence for cloning. In contrast,
screening a cDNA library is laborious, even presuming that a suitable cDNA library is already available
and does not have to be constructed for the purpose.
Quite apart from the labour involved, a cDNA library
may not yield a cDNA clone with a full-length coding region, because, as described above, generating
a full-length cDNA clone may be technically challenging, particularly with respect to long mRNAs.
Furthermore, the sought-after cDNA may be very
rare even in specialized libraries. Does this mean
POGC06 9/11/2001 11:08 AM Page 101
Cloning strategies
that cDNA libraries have been superseded? Despite
the advantages of RT-PCR, there are still reasons for
constructing cDNA libraries. The first reflects the
availability of starting material and the permanence
of the library. A sought-after mRNA may occur in a
source that is not readily available, perhaps a small
number of cells in a particular human tissue. A goodquality cDNA library has only to be constructed
once from this tissue to give a virtually infinite
resource for future use. The specialized library is permanently available for screening. Indeed, the library
may be used as a source from which a specific cDNA
can be obtained by PCR amplification. The second
reason concerns screening strategies. The PCR-based
approaches are dependent upon specific primers.
However, with cDNA libraries, screening strategies
are possible that are based upon expression, e.g.
immunochemical screening, rather than nucleic
acid hybridization (see below).
As discussed above for genomic libraries, PCR can
be used to provide the DNA for library construction when the source is unsuitable for conventional
approaches, e.g. a very small amount of starting
material or fixed tissue. Instead of gene-specific
primers, universal primers can be used that lead to
the amplification of all mRNAs, which can then be
subcloned into suitable vectors. A disadvantage of
PCR-based strategies for cDNA library construction
is that the DNA polymerases used for PCR are more
error-prone than those used conventionally for
second-strand synthesis, so the library may contain
a large number of mutations. There is also likely to
be a certain amount of distortion due to competition
among templates, and a bias towards shorter cDNAs.
A potential problem with RT-PCR is false results
resulting from the amplification of contaminating
genomic sequences in the RNA preparation. Even
trace amounts of genomic DNA may be amplified.
In the study of eukaryotic mRNAs, it is therefore
desirable to choose primers that anneal in different
exons, such that the products expected from the
amplification of cDNA and genomic DNA would be
different sizes or, if the intron is suitably large, so
that genomic DNA would not be amplified at all.
Where this is not possible (e.g. when bacterial RNA
is used as the template), the RNA can be treated with
DNase prior to amplification to destroy any contaminating DNA.
101
Rapid amplification of cDNA ends (RACE)
Another way to address the problem of incomplete
cDNA sequences in libraries is to use a PCR-based
technique for the rapid amplification of cDNA ends
(RACE) (Frohman et al. 1988). Both 5′ RACE and 3′
RACE protocols are available, although 3′ RACE is
usually only required if cDNAs have been generated
using random primers. In each case, only limited
knowledge of the mRNA sequence is required. A
single stretch of sequence within the mRNA is sufficient, so an incomplete clone from a cDNA library
is a good starting-point. From this sequence, specific
primers are chosen which face outwards and which
produce overlapping cDNA fragments. In the two
RACE protocols, extension of the cDNAs from the
ends of the transcript to the specific primers is
accomplished by using primers that hybridize either
at the natural 3′ poly(A) tail of the mRNA, or at a
synthetic poly(dA) tract added to the 5′ end of the
first-strand cDNA (Fig. 6.11). Finally, after amplification, the overlapping RACE products can be
combined if desired, to produce an intact full-length
cDNA.
Although simple in principle, RACE suffers from
the same limitations that affect conventional cDNA
cloning procedures. In 5′ RACE, for example, the
reverse transcriptase may not, in many cases, reach
the authentic 5′ end of the mRNA, but all first-strand
cDNAs, whether full length or truncated, are tailed
in the subsequent reaction, leading to the amplification of a population of variable-length products.
Furthermore, as might be anticipated, since only a
single specific primer is used in each of the RACE
protocols, the specificity of amplification may not be
very high. This is especially problematical where the
specific primer is degenerate. In order to overcome
this problem, a modification of the RACE method has
been devised which is based on using nested primers
to increase specificity (Frohman & Martin 1989).
Strategies for improving the specificity of RACE have
been reviewed (Schaefer 1995, Chen 1996).
Screening strategies
The identification of a specific clone from a DNA
library can be carried out by exploiting either the
sequence of the clone or the structure/function of its
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CHAPTER 6
Race: 5‘ end
Race: 3‘ end
mRNA
cDNA
AAAAAAAAAA
mRNA
T T T T **
**
cDNA
(−) strand
PCR
Denature
Anneal primer
Extend
AAAAAAAAAA
(–) strand
Remove excess 5RT primer
Tail cDNAs with dATP
3‘ amp
AAAAAAAAAA
3‘ amp
5RT
(–) strand
5RT
(+) strand
PCR
(–) strand
Denature
Anneal primer **** T T T T
Extend
T T T T ****
Denature
Anneal primer ****
Extend
***
* TTTT
(+) strand
AAAAAAAAAA
3‘ amp
(+) strand
TR (–) strand
(–) strand
Denature
Anneal primer
Extend
****
5RT
5‘ amp
Denature
Anneal primers:
Up to ~106
copies of cDNA
TR (+) strand
**** T T T T
3‘ amp and ****
Extend
TR (–) strand
3‘ amp
Denature
Anneal primer ****
Extend
(+) strand
TR (–) strand
5‘ amp
****
****
TR (+) strand
TR (–) strand
3‘ amp
Denature
Anneal primers:
**** and 5‘ amp
Extend
****
Up to ~106
copies of cDNA
TR (+) strand
TR (–) strand
5‘ amp
Fig. 6.11 Rapid amplification of cDNA ends (RACE) (Frohman et al. 1988). 3′ Protocol. The mRNA is reverse transcribed using an
oligo(dT17) primer which has a 17 nucleotide extension at its 5′-end. This extension, the anchor sequence, is designed to contain
restriction sites for subsequent cloning. Amplification is performed using the anchor 17-mer (which has a Tm higher than
oligo(dT17)) and a primer specific for the sought-after cDNA. 5′ Protocol. The mRNA is reverse transcribed from a specific primer.
The resultant cDNA is then extended by terminal transferase to create a poly(dA) tail at the 3′-end of the cDNA. Amplification is
performed with the oligo(dT17)/anchor system as used for the 3′ protocol, and the specific primer. Open boxes represent DNA
strands being synthesized; coloured boxes represent DNA from a previous step. The diagram is simplified to show only how the
new product from a previous step is used. Molecules designated TR, truncated, are shorter than full-length (+) or (−) strands.
POGC06 9/11/2001 11:08 AM Page 103
Cloning strategies
103
Box 6.2 Expressed sequence tags (ESTs) for high-throughput genome research
The gold standard in the analysis of individual
genes is a full-length cDNA clone that has been
independently sequenced several times to ensure
accuracy. Such clones are desirable for accurate
archiving and for the detailed mapping of
genomic transcription units, i.e. to determine
the transcriptional start and stop sites, and all
intron/exon boundaries. However, as discussed
in the main text, such clones can be difficult and
expensive to obtain. Technology has not yet
advanced to the stage where full-length cDNAs can
be produced and sequenced in a high-throughput
manner.
Fortunately, full-length cDNA clones are not
required for many types of analysis. Even short cDNA
sequence fragments can be used to unambiguously
identify specific genes and therefore map them
on to physical gene maps or provide information
about their expression patterns. The development
of high-throughput sequencing technology has
allowed thousands of clones to be picked randomly
from cDNA libraries and subjected to single-pass
sequencing to generate 200–300 bp cDNA
signatures called expressed sequence tags
(ESTs) (Wilcox et al. 1991, Okubo et al. 1992).
Although short and somewhat inaccurate, very large
numbers of sequences can be collected rapidly and
inexpensively and deposited into public databases
that can be searched using the Internet. The vast
majority of database sequences are now ESTs rather
than full cDNA or genomic clones. ESTs have been
used for gene discovery, as physical markers on
expressed product. The former applies to any type of
library, genomic or cDNA, and can involve either
nucleic acid hybridization or the PCR. In each case,
the design of the probe or primers can be used to
home in on one specific clone or a group of structurally related clones. Note that PCR screening can
also be used to isolate DNA sequences from uncloned
genomic DNA and cDNA. Screening the product of
a clone applies only to expression libraries, i.e.
libraries where the DNA fragment is expressed to
yield a protein. In this case, the clone can be
genomic maps and for the identification of genes
in genomic clones (e.g. Adams et al. 1991, 1992,
Banfi et al. 1996). Over two million ESTs from
numerous species are currently searchable using the
major public EST database, dbEST (Boguski et al.
1993). The development of EST informatics has been
reviewed (Boguski 1995, Gerhold & Caskey 1996,
Hartl 1996, Okubo & Matsubara 1997). The advent
of ESTs prompted a wide public debate on the
concept of patenting genes. The National Institutes
of Health attempted to patent more than 1000 of
the first EST sequences to be generated, but the
patent application was rejected, predominantly on
the grounds that ESTs are incomplete sequences and
lack precise functional applications (for discussion,
see Roberts 1992).
As well as their use for mapping, ESTs are useful
for expression analysis. PCR primers designed
around ESTs have been used to generate large
numbers of target sequences for cDNA microarrays
(see Box 6.5), and the partial cDNA fragments
used for techniques such as differential-display
PCR are also essentially ESTs (p. 116). The ultimate
EST approach to expression analysis is serial
analysis of gene expression (SAGE). In this technique,
the size of the sequence tag is only 9–10 bp
(the minimum that is sufficient to uniquely identify
specific transcripts) and multiple tags are ligated
into a large concatemer allowing expressed genes
to be ‘read’ by cloning and sequencing the tags
serially arranged in each clone (for details see
Velculescu et al. 1995).
identified because its product is recognized by an
antibody or a ligand of some nature, or because the
biological activity of the protein is preserved and can
be assayed in an appropriate test system.
Sequence-dependent screening
Screening by hybridization
Nucleic acid hybridization is the most commonly
used method of library screening because it is rapid,
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CHAPTER 6
Replica plate onto
nitrocellulose disc
placed on agar in
Petri plate
Incubate
Transformant colonies growing
on agar surface
Retain master
plate
Nitrocellulose disc
removed
Reference
set of
colonies
Pick
positive
colony
(1) Lyse bacteria with 0.5N NaOH
(2) Neutralize
(3) Proteinase
(4) Wash
(5) Bake at 80°C
(1) Hybridize with
32
P-labelled probe
(2) Autoradiography
DNA print
Autoradiograph
it can be applied to very large numbers of clones and,
in the case of cDNA libraries, can be used to identify
clones that are not full-length (and therefore cannot
be expressed).
Grunstein and Hogness (1975) developed a
screening procedure to detect DNA sequences in
transformed colonies by hybridization in situ with
radioactive RNA probes. Their procedure can rapidly
determine which colony among thousands contains
the target sequence. A modification of the method
allows screening of colonies plated at a very high
density (Hanahan & Meselson 1980). The colonies
to be screened are first replica-plated on to a nitrocellulose filter disc that has been placed on the surface
of an agar plate prior to inoculation (Fig. 6.12). A
reference set of these colonies on the master plate is
retained. The filter bearing the colonies is removed
and treated with alkali so that the bacterial colonies
are lysed and the DNA they contain is denatured.
Fig. 6.12 Grunstein–Hogness method for
detection of recombinant clones by colony
hybridization.
The filter is then treated with proteinase K to remove
protein and leave denatured DNA bound to the
nitrocellulose, for which it has a high affinity, in
the form of a ‘DNA print’ of the colonies. The DNA
is fixed firmly by baking the filter at 80°C. The
defining, labelled RNA is hybridized to this DNA
and the result of this hybridization is monitored by
autoradiography. A colony whose DNA print gives a
positive autoradiographic result can then be picked
from the reference plate.
Variations of this procedure can be applied to
phage plaques ( Jones & Murray 1975, Kramer et al.
1976). Benton and Davis (1977) devised a method
called plaque lift, in which the nitrocellulose filter is
applied to the upper surface of agar plates, making
direct contact between plaques and filter. The plaques
contain phage particles, as well as a considerable
amount of unpackaged recombinant DNA. Both
phage and unpackaged DNA bind to the filter and
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Cloning strategies
105
Plate up to 5 104 recombinant
phage on 9 cm square Petri dish
Incubate for 6 –8 h (small plaques),
or overnight (if larger plaques desired)
Cool at 4°C for 1h to stiffen top agar
or top agarose
Nitrocellulose sheet
Overlay plaques with nitrocellulose
sheet for 30 sec to 2 min
Make reference marks for orientation
of sheet with respect to plate
Lift off sheet carefully
Retain plate
Store at 4°C
Phage particles and
recombinant phage DNA
from plaques bind to
nitrocellulose
Fig. 6.13 Benton and Davis’ plaque-lift
procedure.
(1) Place sheet on filter paper soaked
in alkali to denature DNA
(2) Neutralize on filter paper soaked
in neutral buffer
(3) Bake at 80°C in vacuo
(4) Hybridize with labelled nucleic
acid probe
(5) Wash, autoradiograph or
otherwise detect label
can be denatured, fixed and hybridized. This method
has the advantage that several identical DNA prints
can easily be made from a single-phage plate: this
allows the screening to be performed in duplicate,
and hence with increased reliability, and also allows
a single set of recombinants to be screened with two
or more probes. The Benton and Davis (1977) procedure is probably the most widely applied method of
library screening, successfully applied in thousands
of laboratories to the isolation of recombinant phage
by nucleic acid hybridization (Fig. 6.13). More recently, however, library presentation and screening
have become increasingly automated. Box 6.3 considers the advantages of gridded reference libraries.
In place of RNA probes, DNA or synthetic oligonucleotide probes can be used. A number of alternative
labelling methods are also available that avoid the use
Autoradiographic
images of
positive plaques
Pick plugs of agar from retained
plate at positions corresponding
to positive plaques
Isolate recombinant phage
In primary screen of densely-plated
phage library single plaques will not
be identifiable; therefore pick area,
dilute, repeat
of radioactivity. These methods involve the incorporation of chemical labels into the probe, such as digoxigenin or biotin, which can be detected with a specific
antibody or the ligand streptavidin, respectively.
Probe design
A great advantage of hybridization for library screening is that it is extremely versatile. Conditions can be
used in which hybridization is very stringent, so that
only sequences identical to the probe are identified.
This is necessary, for example, to identify genomic
clones corresponding to a specific cDNA or to identify
overlapping clones in a chromosome walk (see below).
Alternatively, less stringent conditions can be used
to identify both identical and related sequences.
This is appropriate where a probe from one species
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CHAPTER 6
Box 6.3 Gridded (arrayed) hybridization reference libraries
Traditionally, library screening by hybridization
involves taking a plaque lift or colony blot, which
generates a replica of the distribution of clones on an
agar plate. However, an alternative is to individually
pick clones and arrange them on the membrane in
the form of a regular grid. Once a laborious process,
gridding or arraying has been considerably simplified
through the use of robotics. Machines can be
programmed to pick clones from microtitre dishes
and spot them onto membranes at a high density;
then the membrane can be hybridized with a
radioactive probe as normal. Using traditional
libraries, positive clones are detected by
autoradiography and the X-ray film must be aligned
with the original plates in order to identify the
corresponding plaques. With gridded libraries,
however, positive signals can be used to obtain sets
of coordinates, which then identify the corresponding
clone from the original microtitre dishes. Since
identical sets of membranes can be easily prepared,
is being used to isolate a homologous clone from
another species (e.g. see Old et al. 1982). Probes corresponding to a conserved functional domain of a
gene may also cross-hybridize with several different
clones in the same species at lower stringency, and
this can be exploited to identify members of a gene
family. The identification of the vertebrate Hox genes
provides an example in which cross-species hybridization was used to identify a family of related clones
(Levine et al. 1984). In this case a DNA sequence
was identified that was conserved between the
Drosophila developmental genes fushi tarazu and
Antennapedia. When this sequence, the homoeobox,
was used to screen a Southern blot of DNA from
other species, including frogs and humans, several
hybridizing bands were revealed. This led to the
isolation of a number of clones from vertebrate cDNA
libraries representing the large family of Hox genes
that play a central role in animal development.
Hybridization thus has the potential to isolate any
sequence from any library if a probe is available. If a
suitable DNA or RNA probe cannot be obtained from
an existing cloned DNA, an alternative strategy is to
duplicates can be distributed to other laboratories
for screening. These laboratories can then determine
the coordinates of their positive signals and order
the corresponding clone from the source laboratory.
Thus, one library can serve a number of different
users and all data can be centralized (Zehetner &
Lehrach 1994). Gridded libraries, while convenient
for screening and data sharing, are more expensive
to prepare than traditional libraries. Therefore,
they are often prepared for high-value libraries with
wide applications, such as genomic libraries cloned
in high-capacity P1, BAC or YAC vectors (Bentley
et al. 1992) and also for valuable cDNA libraries
(Lennon & Lehrach 1991). It is possible to plate
libraries at a density of one clone per well, although
for PACs and BACs it is more common to pool clones
in a hierarchical manner, so that individual clones
may be identified by successive rounds of screening
on smaller subpools (e.g. Shepherd et al. 1994,
Shepherd & Smoller 1994).
make an oligonucleotide probe by chemical synthesis. This requires some knowledge of the amino acid
sequence of the protein encoded by the target clone.
However, since the genetic code is degenerate (i.e.
most amino acids are specified by more than one
codon), degeneracy must be incorporated into probe
design so that a mixture of probes is made, at least
one variant of which will specifically match the target clone. Amino acid sequences known to include
methionine and tryptophan are particularly valuable because these amino acids are each specified by
a single codon, hence reducing the degeneracy of the
resulting probe. Thus, for example, the oligopeptide
His-Phe-Pro-Phe-Met may be identified and chosen
to provide a probe sequence, in which case 32 different oligonucleotides would be required:
T
T T
T
5′ CA TT CCCTT ATG
C C
C
A
G
3′
These 32 different sequences do not have to be synthesized individually because it is possible to perform
POGC06 9/11/2001 11:08 AM Page 107
Cloning strategies
a mixed addition reaction for each polymerization
step. This mixture is then end-labelled with a single
isotopic or alternatively labelled nucleotide, using
an exchange reaction. This mixed-probe method
was originally devised by Wallace and co-workers
(Suggs et al. 1981). To cover all codon possibilities,
degeneracies of 64-fold (Orkin et al. 1983) or even
256-fold (Bell et al. 1984) have been employed successfully. What length of oligonucleotide is required
for reliable hybridization? Even though 11-mers can
be adequate for Southern blot hybridization (SingerSam et al. 1983), longer probes are necessary for
good colony and plaque hybridization. Mixed probes
of 14 nucleotides have been successful, although
16-mers are typical (Singer-Sam et al. 1983).
An alternative strategy is to use a single longer
probe of 40–60 nucleotides. Here the uncertainty at
each codon is largely ignored and instead increased
probe length confers specificity. Such probes are
usually designed to incorporate the most commonly
used codons in the target species, and they may
include the non-standard base inosine at positions of
high uncertainty because this can pair with all four
conventional bases. Such probes are sometimes
termed guessmers. Hybridization is carried out under
low stringency to allow for the presence of mismatches. This strategy is examined theoretically by
Lathe (1985) and has been applied to sequences coding for human coagulation factor VIII (Toole et al.
1984, Wood et al. 1984) and the human insulin
receptor (Ullrich et al. 1985).
Chromosome walking
Earlier in this chapter, we discussed the advantages
of making genomic libraries from random DNA fragments. One of these advantages is that the resulting
fragments overlap, which allows genes to be cloned
by chromosome walking. The principle of chromosome
walking is that overlapping clones will hybridize to
each other, allowing them to be assembled into a
contiguous sequence. This can be used to isolate
genes whose function is unknown but whose genetic
location is known, a technique known as positional
cloning.
To begin a chromosome walk, it is necessary to
have in hand a genomic clone that is known to lie
very close to the suspected location of the target
107
gene. In humans, for example, this could be a restriction fragment length polymorphism that has been
genetically mapped to the same region. This clone is
then used to screen a genomic library by hybridization, which should reveal any overlapping clones.
These overlapping clones are then isolated, labelled
and used in a second round of screening to identify
further overlapping clones, and the process is repeated
to build up a contiguous map. If the same library
is used for each round of screening, previously
identified clones can be distinguished from new
ones, so that walking back and forth along the same
section of DNA is prevented. Furthermore, modern
vectors, such as λDASH and λFIX, allow probes to be
generated from the end-points of a given genomic
clone by in vitro transcription (see Fig. 6.4), which
makes it possible to walk specifically in one direction. In Drosophila, the progress of a walk can also be
monitored by using such probes for in situ hybridization against polytene chromosomes. Monitoring is
necessary due to the dangers posed by repetitive
DNA. Certain DNA sequences are highly repetitive
and are dispersed throughout the genome. Hybridization with such a sequence could disrupt the
orderly progress of a walk, in the worst cases
causing a ‘warp’ to another chromosome. The
probe used for stepping from one genomic clone
to the next must be a unique sequence clone, or a
subclone that has been shown to contain only a
unique sequence.
Chromosome walking is simple in principle, but
technically demanding. For large distances, it is
advisable to use libraries based on high-capacity
vectors, such as BACs and YACs, to reduce the
number of steps involved. Before such libraries were
available, some ingenious strategies were used to
reduce the number of steps needed in a walk. In one
of the first applications of this technology, Hogness
and his co-workers (Bender et al. 1983) cloned
DNA from the Ace and rosy loci and the homoeotic
Bithorax gene complex in Drosophila. The number of
steps was minimized by exploiting the numerous
strains carrying well-characterized inversions and
translocations of specific chromosome regions. A
different strategy, called chromosome jumping, has
been used for human DNA (Collins & Weissman
1984, Poustka & Lehrach 1986). This involves the
circularization of very large genomic fragments
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CHAPTER 6
Box 6.4 A landmark publication. Identification of the cystic
fibrosis gene by chromosome walking and jumping
Cystic fibrosis (CF) is a relatively common severe
autosomal recessive disorder. Until the CF gene was
cloned, there was little definite information about the
primary genetic defect. The cloning of the CF gene
was a breakthrough for studying the biochemistry of
the disorder (abnormal chloride-channel function),
for providing probes for prenatal diagnosis and for
potential treatment by somatic gene therapy or other
means. The publication is especially notable for the
generality of the cloning strategy. In the absence of
any direct functional information about the CF gene,
the chromosomal location of the gene was used as
the basis of the cloning strategy. Starting from
markers identified by linkage analysis as being close
to the CF locus on chromosome 7, a total of about
500 kb was encompassed by a combination of
chromosome walking and jumping. Jumping was
found to be very important to overcome problems
caused by ‘unclonable’ regions which halted the
sequential walks, and in one case achieved a distance
of 100 kb (Collins et al. 1987). In this work, large
numbers of clones were involved, obtained from
several different phage and cosmid genomic libraries.
Among these libraries, one was prepared using the
Maniatis strategy using the lCharon 4A vector, and
several were prepared using the lDASH and lFIX
vectors (Fig. 6.4) after partial digestion of human
genomic DNA with Sau3AI. Cloned regions were
aligned with a map of the genome in the CF region,
obtained by long-range restriction mapping using
rare-cutting enzymes, such as NotI, in combination
with pulsed-field gel electrophoresis (p. 10). The
actual CF gene was detected in this cloned region
by a number of criteria, such as the identification
of open reading frames, the detection of cDNAs
hybridizing to the genomic clones, the detection
of cross-hybridizing sequences in other species and
the presence of CpG islands, which are known
to be associated with the 5′ ends of many genes
in mammals.
From: Rommens et al. (1989) Science 245: 1059–65.
generated by digestion with endonucleases, such
as NotI, which cut at very rare target sites. This
is followed by subcloning of the region covering
the closure of the fragment, thus bringing together
sequences that were located a considerable distance
apart. In this way a jumping library is constructed,
which can be used for long-distance chromosome
walks (Collins et al. 1987, Richards et al. 1988). The
application of chromosome walking and jumping to
the cloning of the human cystic fibrosis gene is discussed in Box 6.4.
but only if there is sufficient information about its
sequence to make suitable primers.*
To isolate a specific clone, PCR is carried out with
gene-specific primers that flank a unique sequence
in the target. A typical strategy for library screening
by PCR is demonstrated by Takumi and Lodish
(1994). Instead of plating the library out on agar, as
would be necessary for screening by hybridization,
pools of clones are maintained in multiwell plates.
Each well is screened by PCR and positive wells are
identified. The clones in each positive well are then
Screening by PCR
* Note that, in certain situations, clever experimental design
can allow the PCR to be used to isolate specific but unknown
DNA sequences. One example of this is 5′ RACE, which is discussed on p. 101. Another is inverse PCR (p. 267), which can
be used to isolate unknown flanking DNA surrounding the
insertion site of an integrating vector. In each case, primers
are designed to bind to known sequences that are joined to the
DNA fragment of interest, e.g. synthetic homopolymer tails,
linkers or parts of the cloning vector.
The PCR is widely used to isolate specific DNA
sequences from uncloned genomic DNA or cDNA,
but it also a useful technique for library screening
(Takumi, 1997). As a screening method, PCR has
the same versatility as hybridization, and the same
limitations. It is possible to identify any clone by PCR
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Cloning strategies
diluted into a series in a secondary set of plates and
screened again. The process is repeated until wells
carrying homogeneous clones corresponding to the
gene of interest have been identified.
There are also several applications where the use
of degenerate primers is favourable. A degenerate
primer is a mixture of primers, all of similar sequence
but with variations at one or more positions. This is
analogous to the use of degenerate oligonucleotides
as hybridization probes, and the primers are synthesized in the same way. A common circumstance
requiring the use of degenerate primers is when the
primer sequences have to be deduced from amino
acid sequences (Lee et al. 1988). Degenerate primers
may also be employed to search for novel members
of a known family of genes (Wilks 1989) or to search
for homologous genes between species (Nunberg
et al. 1989). As with oligonucleotide probes, the
selection of amino acids with low codon degeneracy
is desirable. However, a 128-fold degeneracy in each
primer can be successful in amplifying a single-copy
target from the human genome (Girgis et al. 1988).
Under such circumstances, the concentration of
any individual primer sequence is very low, so mismatching between primer and template must occur
under the annealing conditions chosen. Since mismatching of the 3′-terminal nucleotide of the primer
may prevent efficient extension, degeneracy at this
position is to be avoided.
Screening expression libraries
(expression cloning)
If a DNA library is established using expression
vectors (see Chapter 5), each individual clone can be
expressed to yield a polypeptide. While all libraries
can be screened by hybridization or PCR, as discussed above, expression libraries are useful because
they allow a range of alternative techniques to be
employed, each of which exploits some structural or
functional property of the gene product. This can be
important in cases where the DNA sequence of the
target clone is completely unknown and there is no
strategy available to design a suitable probe or set of
primers.
For higher eukaryotes, all expression libraries are
cDNA libraries, since these lack introns and the
clones are in most cases of a reasonable size. Gener-
109
ally, a random primer method is used for cDNA
synthesis, so there is a greater representation of 5′
sequences. As discussed above, such libraries are
representative of their source, so certain cDNAs are
abundant and others rare. However, it should be
noted that bacterial expression libraries and many
yeast expression libraries are usually genomic, since
there are few introns in bacteria and some yeasts
and very little intergenic DNA. Efficient expression
libraries can be generated by cloning randomly
sheared genomic DNA or partially digested DNA,
and therefore all genes are represented at the same
frequency (Young et al. 1985). A potential problem
with such libraries is that clones corresponding to a
specific gene may carry termination sequences from
the gene lying immediately upstream, which can prevent efficient expression. For this reason, conditions
are imposed so that the size of the fragments for cloning is smaller than that of the target gene, and enough
recombinants are generated for there to be a reasonable chance that each gene fragment will be cloned
in all six possible reading frames (three in each orientation). Considerations for cloning DNA in E. coli expression vectors are discussed further in Chapter 8.
Immunological screening
Immunological screening involves the use of antibodies that specifically recognize antigenic determinants on the polypeptide synthesized by a target
clone. This is one of the most versatile expressioncloning strategies, because it can be applied to any
protein for which an antibody is available. Unlike
the screening strategies discussed below, there is
also no need for that protein to be functional. The
molecular target for recognition is generally an
epitope, a short sequence of amino acids that folds
into a particular three-dimensional conformation
on the surface of the protein. Epitopes can fold
independently of the rest of the protein and therefore often form even when the polypeptide chain
is incomplete or when expressed as a fusion with
another protein. Importantly, many epitopes can
form under denaturing conditions, when the overall
conformation of the protein is abnormal.
The first immunological screening techniques
were developed in the late 1970s, when expression
libraries were generally constructed using plasmid
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CHAPTER 6
Polyvinyl sheet
IgG molecule
Antigen
125
l-labelled lgG
molecule
Fig. 6.14 Antigen–antibody complex formation in the
immunochemical detection method of Broome and Gilbert.
(See text for details.)
vectors. The method of Broome and Gilbert (1978)
was widely used at the time. This method exploited
the fact that antibodies adsorb very strongly to
certain types of plastic, such as polyvinyl, and that
IgG antibodies can be readily labelled with 125I by
iodination in vitro. As usual, transformed cells were
plated out on Petri dishes and allowed to form
colonies. In order to release the antigen from positive
clones, the colonies were lysed, e.g. using chloroform vapour or by spraying with an aerosol of virulent phage (a replica plate is required because this
procedure kills the bacteria). A sheet of polyvinyl
that had been coated with the appropriate antibody
was then applied to the surface of the plate, allowing
antigen–antibody complexes to form. The sheet
was then removed and exposed to 125I-labelled IgG
specific to a different determinant on the surface of
the antigen (i.e. a determinant not involved in the
initial binding of the antigen to the antibody-coated
sheet (Fig. 6.14)). The sheet was then washed and
exposed to X-ray film. The clones identified by this
procedure could then be isolated from the replica
plate. Note that this ‘sandwich’ technique is applicable only where two antibodies recognizing different
determinants of the same protein are available. However, if the protein is expressed as a fusion, antibodies
that bind to each component of the fusion can be
used, efficiently selecting for recombinant molecules.
While plasmid libraries have been useful for
expression screening (Helfman et al. 1983, Helfman
& Hughes 1987), it is much more convenient to use
bacteriophage-λ insertion vectors, because these
have a higher capacity and the efficiency of in vitro
packaging allows large numbers of recombinants to
be prepared and screened. Immunological screening
with phage-λ cDNA libraries was introduced by
Young and Davies (1983) using the expression
vector λgt11, which generates fusion proteins with
β-galactosidase under the control of the lac promoter
(see Box 6.1 for a discussion of λgt11 and similar
fusion vectors, such as λZAP). In the original technique, screening was carried out using colonies of
induced lysogenic bacteria, which required the production of replica plates, as above. A simplification of
the method is possible by directly screening plaques
of recombinant phage. In this procedure (Fig. 6.15),
the library is plated out at moderately high density
(up to 5 × 104 plaques/9 cm2 plate), with E. coli
strain Y1090 as the host. This E. coli strain overproduces the lac repressor and ensures that no expression of cloned sequences (which may be deleterious
to the host) takes place until the inducer isopropyl-β-thiogalactoside (IPTG) is presented to the infected
cells. Y1090 is also deficient in the lon protease,
hence increasing the stability of recombinant fusion
proteins. Fusion proteins expressed in plaques are
absorbed on to a nitrocellulose membrane overlay,
and this membrane is processed for antibody screening. When a positive signal is identified on the
membrane, the positive plaque can be picked from
the original agar plate (a replica is not necessary)
and the recombinant phage can be isolated.
The original detection method using iodinated
antibodies has been superseded by more convenient
methods using non-isotopic labels, which are also
more sensitive and have a lower background of nonspecific signal. Generally, these involve the use of
unlabelled primary antibodies directed against the
polypeptide of interest, which are in turn recognized
by secondary antibodies carrying an enzymatic label.
As well as eliminating the need for isotopes, such
methods also incorporate an amplification step,
since two or more secondary antibodies bind to the
primary antibody. Typically, the secondary antibody
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Cloning strategies
lacZ
cl857
Sam
cl857
Sam
λgt11
111
Insert cDNA at EcoRI site of λgt11 or
in polylinker region of λZAP
EcoRI
lacZ
λZAP
Polylinker
Plate out library on E. coli Y1090
Incubate for 4–6 h at 37°C to obtain
small plaques
Nitrocellulose sheet
Overlay plaques with nitrocellulose
sheet previously soaked in IPTG;
this induces expression from lac
promoter
Incubate for 4 hours
Carefully remove nitrocellulose sheet,
which will have adsorbed fusion
proteins expressed in recombinant
phage plaques, i.e. plaque-lift
Retain plate
Pick positive plaque from retained
plaque
Fig. 6.15 Immunochemical screening
of λgt11 or λZAP recombinant
plaques.
recognizes the species-specific constant region of the
primary antibody and is conjugated to either horseradish peroxidase (De Wet et al. 1984) or alkaline
phosphatase (Mierendorf et al. 1987), each of which
can in turn be detected using a simple colorimetric
assay carried out directly on the nitrocellulose filter.
Polyclonal antibodies, which recognize many different epitopes, provide a very sensitive probe for
immunological screening, although they may also
cross-react to proteins in the expression host. Monoclonal antibodies and cloned antibody fragments
can also be used, although the sensitivity of such
reagents is reduced because only a single epitope is
recognized.
Screen nitrocellulose plaque-lift
with specific antibody to detect
fusion protein
Identify positive plaque
South-western and north-western screening
We have seen how fusion proteins expressed in
plaques produced by recombinant λgt11 or λZAP
vectors may be detected by immunochemical screening. A closely related approach has been used for the
screening and isolation of clones expressing sequencespecific DNA-binding proteins. As above, a plaque
lift is carried out to transfer a print of the library on to
nitrocellulose membranes. However, the screening
is carried out, without using an antibody, by incubating the membranes with a radiolabelled doublestranded DNA oligonucleotide probe, containing the
recognition sequence for the target DNA-binding
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CHAPTER 6
protein. This technique is called south-western screening, because it combines the principles of Southern
and western blots. It has been particularly successful
in the isolation of clones expressing cDNA sequences
corresponding to certain mammalian transcription
factors (Singh et al. 1988, Staudt et al. 1988, Vinson
et al. 1988, Katagiri et al. 1989, Williams et al. 1991,
Xiao et al. 1991). A limitation of this technique is that,
since individual plaques contain only single cDNA
clones, transcription factors that function only in the
form of heterodimers or as part of a multimeric complex do not recognize the DNA probe and the corresponding cDNAs cannot be isolated. Clearly the
procedure can also be successful only in cases where
the transcription factor remains functional when
expressed as a fusion polypeptide. It is also clear that
the affinity of the polypeptide for the specific DNA
sequence must be high, and this has led to the preferential isolation of certain types of transcription factor (reviewed by Singh 1993). More recently, a
similar technique has been used to isolate sequencespecific RNA-binding proteins, in this case using a
single-stranded RNA probe. By analogy to the above,
this is termed north-western screening and has been
successful in a number of cases (e.g. see Qian &
Wilusz 1993; reviewed by Bagga & Wilusz 1999).
Both south-western and north-western screening are
most efficient when the oligonucleotide contains the
binding sequence in multimeric form. This may mean
that several fusion polypeptides on the filter bind to
each probe, hence greatly increasing the average
dissociation time. To minimize non-specific binding,
a large excess of unlabelled double-stranded DNA
(or single-stranded RNA) is mixed with the specific
probe. However, it is usually necessary to confirm
the specificity of binding in a second round of screening, using the specific oligonucleotide probe and
one or more alternative probes containing a similar
sequences that are not expected to be recognized.
Screening with alternative ligands
As well as DNA and RNA, a whole range of alternative ‘ligands’ can be used to identify polypeptides that
specifically bind certain molecules (for example, as
an alternative to south-western screening). Such
techniques are not widely used because they generally have a low sensitivity and their success depends
on the preservation of the appropriate interacting
domain of the protein when exposed on the surface
of a nitrocellulose filter. Furthermore, as discussed in
Chapter 9, the yeast two-hybrid system and its
derivatives now provide versatile assay formats for
many specific types of protein–protein interaction,
with the advantage that such interactions are tested
in living cells, so the proteins involved are more
likely to retain their functional interacting domains.
Functional cloning
Finally, we consider screening methods that depend
on the full biological activity of the protein. This is
often termed functional cloning. In contrast to positional cloning, described above, functional cloning
is possible in complete ignorance of the whereabouts
of the gene in the genome and requires no prior
knowledge of the nucleotide sequence of the clone or
the amino acid sequence of its product. As long as
the expressed protein is functional and that function
can be exploited to screen an expression library, the
corresponding clone can be identified.
Screening by functional complementation
Functional complementation is the process by which
a particular DNA sequence compensates for a missing function in a mutant cell, and thus restores the
wild-type phenotype. This can be a very powerful
method of expression cloning, because, if the mutant
cells are non-viable under particular growth conditions, cells carrying the clone of interest can be positively selected, allowing the corresponding clones to
be isolated.
Ratzkin and Carbon (1977) provide an early
example of how certain eukaryotic genes can be
cloned on the basis of their ability to complement
auxotrophic mutations in E. coli. These investigators
inserted fragments of yeast DNA, obtained by mechanical shearing, into the plasmid ColEl, using a
homopolymer-tailing procedure. They transformed
E. coli hisB mutants, which are unable to synthesize
histidine, with the recombinant plasmids and plated
the bacteria on minimal medium. In this way, they
selected for complementation of the mutation and
isolated clones carrying an expressed yeast his gene.
If the function of the gene is highly conserved, it is
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Cloning strategies
113
Wild type mouse
shaker-2 mouse
Genomic DNA
BAC
genomic
library
Homozygous
shaker-2 eggs
Fig. 6.16 Functional
complementation in transgenic
mice to isolate the Shaker-2 gene.
Homozygous shaker-2 fertilized mouse
eggs were injected with BAC clones
derived from the Shaker-2 candidate
region of a wild-type mouse. Progeny
were screened for restoration of the
wild-type phenotype, thus identifying
the BAC clone corresponding to the
Shaker-2 gene. This clone is then
sequenced and used to isolate and map
the corresponding human disease
gene DFNB3.
BAC clones
spanning shaker-2
candidate region
quite possible to carry out functional cloning of, for
example, mammalian proteins in bacteria and yeast.
Thus, complementation in yeast has been used to
isolate cDNAs for a number of mammalian metabolic enzymes (e.g. Botstein & Fink 1988) and certain
highly conserved transcription factors (e.g. Becker
et al. 1991), as well as regulators of meiosis in plants
(Hirayama et al. 1997). This approach can also
be used in mammalian cells, as demonstrated by
Strathdee et al. (1992), who succeeded in isolating
the FACC gene, corresponding to complementation
group C of Fanconi’s anaemia. Generally a pool system is employed, where cells are transfected with a
complex mix of up to 100 000 clones. Pools that
successfully complement the mutant phenotype are
Inject individual
BAC clones
Sequence
Screen human library
Identify clone
that corrected defect
Map of human gene
then subdivided for a further round of transfection,
and the procedure repeated until the individual
cDNA responsible is isolated.
Functional complementation is also possible in
transgenic animals and plants. In this way, Probst
et al. (1998) were able to clone the mouse deafnessassociated gene Shaker-2, and from there its human
homologue, DFNB3 (Fig. 6.16). The shaker-2 mutation had previously been mapped to a region of
the mouse genome that is syntenic to the region
involved in a human deafness disorder. BAC clones
corresponding to this region were therefore prepared
from wild-type mice and microinjected into the eggs
of shaker-2 mutants. The resulting transgenic mice
were screened for restoration of a normal hearing
POGC06 9/11/2001 11:08 AM Page 114
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CHAPTER 6
phenotype, allowing a BAC clone corresponding to
the functional Shaker-2 gene to be identified. The
gene was shown to encode a cytoskeletal myosin
protein. This was then used to screen a human
genomic library, resulting in the identification of the
equivalent human gene. Note that no sequence
information was required for this screening procedure, and without the functional assay there would
have been no way to identify either the mouse or
human gene except through a laborious chromosome walk from a linked marker. The recent development of high-capacity transformation vectors for
plants (p. 236) has allowed similar methods to be
used to identify plant genes (e.g. Sawa et al. 1999,
Kubo & Kakimoto 2001).
Screening by ‘gain of function’
Complementation analysis can be used only if an
appropriate mutant expression host is available. In
many cases, however, the function of the target
gene is too specialized for such a technique to work
in a bacterial or yeast expression host and, even in
a higher eukaryotic system, loss of function in the
host may be fully or partially compensated by one or
more other genes. As an alternative, it may be possible to identify clones on the basis that they confer a
gain of function on the host cell. In some cases, this
gain of function is a selectable phenotype that allows
cells containing the corresponding clone to be positively selected. For example, in an early example
of the expression of a mammalian gene in E. coli,
Chang et al. (1978) constructed a population of
recombinant plasmids containing cDNA derived
from unfractionated mouse mRNA. This population
of mRNA molecules was expected to contain the
transcript for dihydrofolate reductase (DHFR). Mouse
DHFR is much less sensitive to inhibition by the drug
trimethoprim than E. coli DHFR, so growing transformants in medium containing the drug allowed
selection for those cells containing the mouse Dhfr
cDNA.
In other cases, the phenotype conferred by the
clone of interest is not selectable, but can be detected
because it causes a visible change in phenotype. In
mammalian cells, for example, clones corresponding
to cellular oncogenes have been identified on the
basis of their ability to stimulate the proliferation
of quiescent mouse 3T3 fibroblast cells either in
culture or when transplanted into ‘nude mice’ (e.g.
Brady et al. 1985). Many different specific assays
have also been developed for the functional cloning
of cDNAs encoding particular types of gene product.
For example, Xenopus melanophores have been used
for the functional cloning of G-protein-coupled
receptors. Melanophores are dark cells containing
many pigment organelles, called melanosomes. A
useful characteristic of these organelles is that they
disperse when adenyl cyclase or phospholipase C
are active and aggregate when these enzymes are
inhibited. Therefore, the expression of cDNAs encoding G-protein-coupled receptors and many types of
receptor tyrosine kinases leads to redistribution of
pigmentation within the cell, which can be used
as an assay for the identification of receptor cDNAs
(reviewed be Lerner 1994).
Difference cloning
Difference cloning refers to a range of techniques
used to isolate sequences that are represented in one
source of DNA but not another. Normally this
means differentially expressed cDNAs, representing
genes that are active in one tissue but inactive in
another, but the technique can also be applied to
genomic DNA to identify genes corresponding to
deletion mutants. There are a number of cell-based
differential cloning methods and also a range of PCR
techniques. Each method follows one of two principles: either the differences between two sources are
displayed, allowing differentially expressed clones to
be visually identified, or the differences are exploited
to generate a collection of clones that are enriched
for differentially expressed sequences. The analysis
of differential gene expression has taken on new
importance recently with the advent of highthroughput techniques allowing the monitoring of
many and, in some cases, all genes simultaneously.
Difference cloning with DNA libraries
Displaying differences – differential screening
An early approach to difference cloning was differential
screening, a simple variation on normal hybridizationbased library screening protocols that is useful for
POGC06 9/11/2001 11:08 AM Page 115
Cloning strategies
the identification of differentially expressed cDNAs
that are also moderately abundant (e.g. Dworkin
& Dawid 1980). Let us consider, for example, the
isolation of cDNAs derived from mRNAs which are
abundant in the gastrula embryo of the frog Xenopus
but which are absent, or present at low abundance,
in the egg. A cDNA library is prepared from gastrula
mRNA. Replica filters carrying identical sets of
recombinant clones are then prepared. One of these
filters is then probed with 32P-labelled mRNA (or
cDNA) from gastrula embryos and one with 32Plabelled mRNA (or cDNA) from the egg. Some
colonies will give a positive signal with both probes;
these represent cDNAs derived from mRNA types
that are abundant at both stages of development.
Some colonies will not give a positive signal with
either probe; these correspond to mRNA types
present at undetectably low abundance in both
tissues. This is a feature of using complex probes,
which are derived from mRNA populations rather
than single molecules: only abundant or moderately abundant sequences in the probe carry a
significant proportion of the label and are effective
in hybridization. Importantly, some colonies give a
positive signal with the gastrula probe, but not
with the egg probe. These can be visually identified
and should correspond to differentially expressed
sequences.
A recent resurgence in the popularity of differential screening has come about through the development of DNA microarrays (Schena et al. 1995). In
this technique, cDNA clones are transferred to a
miniature solid support in a dense grid pattern and
screened simultaneously with complex probes from
two sources, which are labelled with different fluorophores. Clones that are expressed in both tissues will
fluoresce in a colour that represents a mixture of
fluorophores, while differentially expressed clones
will fluoresce in a colour closer to the pure signal
of one or other of the probes. A similar technique
involves the use of DNA chips containing densely
arrayed oligonucleotides. These methods are compared in Box 6.5.
Enrichment for differences – subtractive cloning
An alternative to differential screening is to generate
a library that is enriched in differentially expressed
115
clones by removing sequences that are common to
two sources. This is called a subtracted cDNA library
and should greatly assist the isolation of rare cDNAs.
If we use the same example as above, the aim of the
experiment would be to generate a library enriched
for cDNAs derived from gastrula-specific mRNAs.
This would be achieved by hybridizing first-strand
cDNAs prepared from gastrula mRNA with a large
excess of mRNA from Xenopus oocytes. If this driver
population is labelled in some way, allowing it to be
removed from the mixed population, only gastrulaspecific cDNAs would remain behind. A suitable
labelling method would be to add biotin to all the
oocyte mRNA, allowing oocyte/gastrula RNA/cDNA
hybrids as well as excess oocyte mRNA to be subtracted by binding to streptavidin, for which biotin
has great affinity (Duguid et al. 1988, Rubinstein
et al. 1990). Highly enriched libraries can be prepared by several rounds of extraction with driver
mRNA, resulting in highly enriched subtracted
libraries (reviewed by Sagerstrom et al. 1997).
An example of subtractive cDNA cloning and
differential screening is provided by Nedivi et al.
(1993). These investigators were interested in the
isolation of rat cDNAs that are induced in a particular
region of the brain (the dentate gyrus (DG)) known
to be involved in learning and memory. The inducing stimulus was kainate, a glutamate analogue
that induces seizures and memory-related synaptic
changes. Poly(A)+ RNA was extracted from the DG
of kainate-treated animals and used for first-strand
cDNA synthesis. Ubiquitous sequences present in
the activated DG cDNA preparation were hybridized
with an excess of poly(A)+ RNA from total uninduced rat brain. This RNA had previously been
biotinylated (using a photobiotinylation procedure)
and so hybrids and excess RNA could be removed
using a streptavidin extraction method (Sive & St
John 1988). The unhybridized cDNA was then converted into double-stranded form by conventional
methods and used to construct the subtracted cDNA
library in λZAP. This subtracted library was differentially screened using radiolabelled cDNA from
activated and non-activated DG as the differential
probes. A large number of activated DG clones were
isolated, of which 52 were partially sequenced. Onethird of these clones corresponded to known genes;
the remainder were new.
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CHAPTER 6
Box 6.5 Differential screening with DNA chips
cDNA microarrays
Miniaturization and automation have facilitated
the development of DNA microarrays, in which
DNA sequences are displayed on the surface of a
small ‘chip’ of either nylon or glass. In the initial
description of this technology (Schena et al. 1995),
up to 10 000 cDNA clones, each in the region of
several hundred nucleotides in length, could be
arrayed on a single microscope slide. The cDNAs
were either obtained from an existing library or
generated de novo by PCR. In each case, the machine
transfers a small amount of liquid from a standard
96-well microtitre plate on to a poly-l-lysine-coated
microscope slide, and the DNA is fixed in position
by UV irradiation. Arrays are used predominantly for
the multiplex analysis of gene expression profiles.
Total RNA is used to prepare fluorescently labelled
cDNA probes and signals are detected using a laser.
Each hybridization experiment generates a large
amount of data. Comparisons of expression profiles
generated using probes from different sources can
identify genes that are differentially expressed in
various cell types, at different developmental stages
or in response to induction (reviewed by Schena
et al. 1998, Xiang & Chen 2000). There have been
many successes with this relatively new technology,
including the identification of genes involved in the
development of the nervous system (Wen et al. 1998)
and genes involved in inflammatory disease (Heller
et al. 1997). Arrays have been constructed including
every gene in the genome of E. coli (Tao et al. 1999)
Difference cloning by PCR
Displaying differences – differential-display
PCR and arbitrarily primed PCR
As expected, PCR-based methods for difference
cloning are more sensitive and rapid than librarybased methods, and can be applied to small amounts
of starting material. Two similar methods have been
and most genes of the yeast Saccharomyces
cerevisiae (De Risi et al. 1997). This has allowed
comprehensive parallel analysis of the expression of
all genes simultaneously in a variety of experimental
assays (Cho et al. 1998, Chu et al. 1998, Spellman
et al. 1998, Jelinsky & Samson 1999). It is likely that
complete genome arrays will be available for higher
eukaryotes, including humans, within the next few
years, offering an unprecedented ability to capture
functional snapshots of the genome in action.
Oligonucleotide chips
An alternative to spotting presynthesized cDNAs or
ESTs on to slides is to synthesize oligonucleotides
in situ on silicon or glass wafers, using similar
processes to those used in the manufacture of
semiconductors (Lockhart et al. 1996, Shalon et al.
1996). Using current techniques, up to 1 000 000
oligonucleotides can by synthesized in tightly
packed regular arrays on chips approximately
1 cm2 (Lipshutz et al. 1999). Unlike cDNA arrays, a
hybridizing probe sequence is recognized not by a
single cognate cDNA, but by a combination of short
oligonucleotides, from which its sequence can be
deduced (reviewed by Southern 1996a,b). Chips
are more versatile than arrays, because they can be
used not only for expression analysis but also for
DNA sequencing (resequencing) (Chee et al. 1996)
and the analysis of differences between genomes
at the level of single nucleotide polymorphisms
(Hacia 1999).
described which use pairs of short arbitrary primers
to amplify pools of partial cDNA sequences. If the
same primer combinations are used to amplify
cDNAs from two different tissues, the products can
be fractionated side by side on a sequencing gel, and
differences in the pattern of bands generated, the
mRNA fingerprint, therefore reveal differentially
expressed genes (Fig. 6.17). Essentially, the distinction between the two techniques concerns the
POGC06 9/11/2001 11:08 AM Page 117
Cloning strategies
117
Box 6.6 A landmark publication. Subtraction cloning of the human
Duchenne muscular dystrophy (DMD) gene
While most subtractive cloning experiments involve
cDNAs, this publication reports one of the few
successful attempts to isolate a gene using a
subtracted genomic library. The study began with
the identification of a young boy, known as ‘BB’,
who suffered from four X-linked disorders, including
DMD. Cytogenetic analysis showed that the boy
had a chromosome deletion in the region Xp21,
which was known to be the DMD locus. Kunkel’s
group then devised a subtraction-cloning procedure
to isolate the DNA sequences that were deleted
in BB. Genomic DNA was isolated from BB and
randomly sheared, generating fragments with
blunt ends and non-specific overhangs. DNA was
also isolated from an aneuploid cell line with four
(normal) X chromosomes. This DNA was digested
with the restriction enzyme MboI, generating sticky
ends suitable for cloning. The MboI fragments were
mixed with a large excess of the randomly sheared
DNA from BB, and the mixture was denatured and
then persuaded to reanneal extensively, using phenol
enhancement. The principle behind the strategy was
primer used for first-strand cDNA synthesis. In the
differential-display PCR technique (Liang & Pardee
1992), the antisense primer is an oligo-dT primer
with a specific two-base extension, which thus
binds at the 3′ end of the mRNA. Conversely, in the
arbitrarily primed PCR method (Welsh et al. 1992),
the antisense primer is arbitrary and can in principle anneal anywhere in the message. In each case,
an arbitrary sense primer is used, allowing the
amplification of partial cDNAs from pools of several
hundred mRNA molecules. Following electrophoresis, differentially expressed cDNAs can be excised
from the gel and characterized further, usually to
confirm its differential expression.
Despite the fact that these display techniques
are problematical and appear to generate a large
number of false-positive results, there have been
remarkable successes. In the original report by
that, since the randomly fragmented DNA was
present in a vast excess, most of the DNA from the
cell line would be sequestered into hybrid DNA
molecules that would be unclonable. However,
those sequences present among the MboI fragments
but absent from BB’s DNA due to the deletion would
only be able to reanneal to complementary strands
from the cell line. Such strands would have intact
MboI sticky ends and could therefore be ligated into
an appropriate cloning vector. Using this strategy,
Kunkel and colleagues generated a genomic library
that was highly enriched for fragments corresponding
to the deletion in BB. Subclones from the library
were tested by hybridization against normal DNA
and DNA from BB to confirm that they mapped
to the deletion. To confirm that the genuine DMD
gene had been isolated, the positive subclones
were then tested against DNA from many other
patients with DMD, revealing similar deletions
in 6.5% of cases.
From Kunkel (1986) Nature 322: 73–77.
Liang and Pardee (1992), the technique was used
to study differences between tumour cells and
normal cells, resulting in the identification of a
number of genes associated with the onset of cancer
(Liang et al. 1992). Further cancer-related gene
products have been discovered by other groups
using differential display (Sager et al. 1993, Okamato
& Beach 1994). The technique has also been used
successfully to identify developmentally regulated
genes (e.g. Adati et al. 1995) and genes that are
induced by hormone treatment (Nitsche et al. 1996).
An advantage of display techniques over subtracted libraries is that changes can be detected in
related mRNAs representing the same gene family.
In subtractive-cloning procedures, such differences
are often overlooked because the excess of driver
DNA can eliminate such sequences (see review by
McClelland et al. 1995).
POGC06 9/11/2001 11:08 AM Page 118
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CHAPTER 6
(a)
A G T T T T T T T T GG T T T T T T T T CG T T T T T T T T T G T T T T T T T T
A A T T T T T T T T GA T T T T T T T T CA T T T T T T T T T A T T T T T T T T
=
NV T T T T T T T T
A C T T T T T T T T GC T T T T T T T T C C T T T T T T T T T C T T T T T T T T
(b)
AAAAAAAA
AAAAAAAA
AAAAAAAA
NV T T T T T T T T
AAAAAAAA
NV T T T T T T T T
NV T T T T T T T T
N9
NV T T T T T T T T
N9
NV T T T T T T T T
AAAAAAAA
NV T T T T T T T T
AAAAAAAA
(c)
Fig. 6.17 Summary of the differential mRNA display technique, after Liang and Pardee (1992). (a) A set of 12 oligo(dT) primers
is synthesized, each with a different two-base extension; the generic designation for this primer set is NVTTTTTTTTT, where N is
any nucleotide and V is any nucleotide except T. (b) Messenger RNA from two sources is then converted into cDNA using these
primers, generating 12 non-overlapping pools of first strand cDNA molecules for each source, The PCR is then carried out using
the appropriate oligo(dT) primer and a set of arbitrary 9-mers (N9), which may anneal anywhere within the cDNA sequence. This
facilitates the amplification of pools of cDNA fragments, essentially the same as expressed sequence tages (ESTs). (c) Pools of PCR
products, derived from alternative mRNA sources but amplified with the same pair of primers, are then compared side by side on a
sequencing gel. Bands present in one lane but absent from the other are likely to represent differentially-expressed genes. The
corresponding bands can be excised from the sequencing gel and the PCR products subcloned, allowing sequence annotation and
expression analysis, e.g. by northern blot or in situ hybridization, to confirm differential expression.
Enrichment for differences – representational
difference analysis
Representational difference analysis is a PCR subtraction technique, i.e. common sequences between
two sources are eliminated prior to amplification.
The method was developed for the comparative
analysis of genomes (Lisitsyn et al. 1993) but has
been modified for cloning differentially expressed
genes (Hubank & Schatz 1994). Essentially, the
technique involves the same principle as subtraction
hybridization, in that a large excess of a DNA from
one source, the driver, is used to make common
sequences in the other source, the tester, unclonable
(in this case unamplifiable). The general scheme is
shown in Fig. 6.18. cDNA is prepared from two
sources, digested with restriction enzymes and
amplified. The amplified products from one source
POGC06 9/11/2001 11:08 AM Page 119
Cloning strategies
mRNA
119
cDNA
Restriction digest
(4 cutter)
Ligate 12/24 linker
Melt 12-mer; fill in
PCR
Representations
Tester
Driver
Digest;
ligate new 12/24 linker
Tester:Driver
Digest
Mix 1 : 100; Melt; Hybridize
Tester:Tester
Driver:Driver
Exponential amplification
Retained
No amplification
Eliminated
Melt 12-mer;
Fill in; PCR
Fig. 6.18 Basic strategy for cDNA-based
representational difference analysis.
See text for details.
Linear amplification
Digest with
Mung Bean Nuclease
PCR
are then annealed to specific linkers that provide
annealing sites for a unique pair of PCR primers.
These linkers are not added to the driver cDNA. A
large excess of driver cDNA is then added to the
tester cDNA and the populations are mixed. Driver/
driver fragments possess no linkers and cannot be
First difference product
amplified, while driver/tester fragments possess only
one primer annealing site and will only be amplified
in a linear fashion. However, cDNAs that are present
only in the tester will possess linkers on both strands
and will be amplified exponentially and can therefore be isolated and cloned.
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CHAPTER 7
Sequencing and mutagenesis
Introduction
DNA sequencing is a fundamental requirement
for modern gene manipulation. Knowledge of the
sequence of a DNA region may be an end in its
own right, perhaps in understanding an inherited
human disorder. More importantly, sequence information is a prerequisite for planning any substantial
manipulation of the DNA; for example, a computer
search of the sequence for all known restrictionendonuclease target sites will provide a complete
and precise restriction map. Similarly, mutants are
an essential prerequisite for any genetic study and
never more so than in the study of gene structure
and function relationships.
Classically, mutants are generated by treating the
test organism with chemical or physical agents that
modify DNA (mutagens). This method of mutagenesis has been extremely successful, as witnessed by
the growth of molecular biology, but suffers from
a number of disadvantages. First, any gene in the
organism can be mutated and the frequency with
which mutants occur in the gene of interest can be
very low. This means that selection strategies have
to be developed. Second, even when mutants with
the desired phenotype are isolated, there is no guarantee that the mutation has occurred in the gene of
interest. Third, prior to the development of gene
cloning and sequencing techniques, there was no
way of knowing where in the gene the mutation had
occurred and whether it arose by a single base
change, an insertion of DNA or a deletion.
As techniques in molecular biology have developed, so that the isolation and study of a single gene
is not just possible but routine, so mutagenesis has
also been refined. Instead of crudely mutagenizing
many cells or organisms and then analysing many
thousands or millions of offspring to isolate a desired
mutant, it is now possible to specifically change any
given base in a cloned DNA sequence. This technique
is known as site-directed mutagenesis. It has become a
basic tool of gene manipulation, for it simplifies DNA
manipulations that in the past required a great
deal of ingenuity and hard work, e.g. the creation or
elimination of cleavage sites for restriction endonucleases. The importance of site-directed mutagenesis
goes beyond gene structure–function relationships,
for the technique enables mutant proteins to be
generated with very specific changes in particular
amino acids (protein engineering). Such mutants
facilitate the study of the mechanisms of catalysis,
substrate specificity, stability, etc (see p. 299 et seq.).
Basic DNA sequencing
The first significant DNA sequence to be obtained
was that of the cohesive ends of phage-λ DNA (Wu &
Taylor 1971), which are only 12 bases long. The
methodology used was derived from RNA sequencing and was not applicable to large-scale DNA
sequencing. An improved method, plus and minus
sequencing, was developed and used to sequence
the 5386 bp phage ΦX 174 genome (Sanger et al.
1977a). This method was superseded in 1977 by
two different methods, that of Maxam and Gilbert
(1977) and the chain-termination or dideoxy method
(Sanger et al. 1977b). For a while the Maxam and
Gilbert (1977) method, which makes use of chemical
reagents to bring about base-specific cleavage of
DNA, was the favoured procedure. However, refinements to the chain-termination method meant that by
the early 1980s it became the preferred procedure.
To date, most large sequences have been determined
using this technology, with the notable exception of
bacteriophage T7 (Dunn & Studier 1983). For this
reason, only the chain-termination method will be
described here.
The chain-terminator or dideoxy procedure for
DNA sequencing capitalizes on two properties of
DNA polymerases: (i) their ability to synthesize faithfully a complementary copy of a single-stranded
DNA template; and (ii) their ability to use 2′,3′-
POGC07 9/11/2001 11:07 AM Page 121
HO
γ
β
α
O
O
O
P
O–
O
P
O–
O
P
Base
(A, G, C or T)
O
O–
5’
CH2
4’
HO
O
H
β
α
O
O
P
O
P
O–
1’
H
γ
O
O
Base
(A, G, C or T)
P
O–
O
O–
4’
H
H
3’
OH
5’
CH2
O
H
H
2’
H
2’
H
3’
OH group missing H
Normal deoxynucleoside triphosphate
(i.e. 2’ deoxynucleotide)
1’
H
H
Dideoxynucleoside triphosphate
(i.e. 2’, 3’ dideoxynucleotide)
Fig. 7.1 Dideoxynucleoside triphosphates act as chain terminators because they lack a 3′ OH group. Numbering of the carbon
atoms of the pentose is shown (primes distinguish these from atoms in the bases). The α, β and γ phosphorus atoms are indicated.
5'
3'
CATACGTGG CCTTACG
Add 5' CGTAAGG3' primer
5'
3'
CATACGTGGCCTTACG
GGAATGC 5'
Add d*ATP, d*CTP, d*GTP and d*TTP
Add
ddATP
+
Klenow
fragment
Add
ddCTP
+
Klenow
fragment
CGTAAGG*C*CdA
and
CGTAAGG*C*C*A*G*G*TdA
Add
ddGTP
+
Klenow
fragment
Add
ddTTP
+
Klenow
fragment
CGTAAGG*C*C*A*CdG
and
CGTAAGG*C*C*A*C*G*T*A*TdG
CGTAAGGdC
and
CGTAAGG*CdC
and
CGTAAGG*C*C*AdC
CGTAAGG*C*C*A*C*GdT
and
CGTAAGG*C*C*A*C*G*T*AdT
Electrophorese and autoradiograph
Sequence
Fig. 7.2 DNA sequencing with
dideoxynucleoside triphosphates as
chain terminators. In this figure
asterisks indicate the presence of 32P
and the prefix ‘d’ indicates the presence
of a dideoxynucleoside. At the top of
the figure the DNA to be sequenced is
enclosed within the box. Note that
unless the primer is also labelled with
a radioisotope the smallest band with
the sequence CGTAAGGdC will not be
detected by autoradiography, as no
labelled bases were incorporated.
A
C
G
T
CGTAAGGCCACGTATdG
CGTAAGGCCACGTAdT
CGTAAGGCCACGTdA
CGTAAGGCCACGdT
CGTAAGGCCACdG
CGTAAGGCCAdC
CGTAAGGCCdA
CGTAAGGCdC
CGTAAGGdC
Direction of
electrophoresis
POGC07 9/11/2001 05:21 PM Page 122
122
CHAPTER 7
Sequencing gel autoradiograph
A
C
G
T
Electrophoresis
T
3‘
C
G
C
A
G
T
C
C
T
A
G
C
T
T
A
G
C
G
G
EcoRI
BamI
SalI
SalI
PstI
BamI
5‘
Fig. 7.3 Enlarged autoradiograph of a
sequencing gel obtained with the chainterminator DNA sequencing method.
EcoRI
5′
3′
ATGACCATGATTACGAATTCCCCGGATCCGTCGACCTGCAGGTCGACGGATCCGGGGAATTCACTGGCCGTCGTTTTACAACGTCGTGACTGGG
TTGCAGCACTGACCC
AccI HincII
1 2 3 4
3′
AccI HincII
Direction of
lac translation
Start of
translation of
lac α fragment
Synthesis of
DNA in
sequencing
reaction
5′
Primer
oligonucleotide
Fig. 7.4 Sequence of M13 mp7 DNA in the vicinity of the multipurpose cloning region. The upper sequence is that of M13 mp7
from the ATG start codon of the β-galactosidase α fragment, through the multipurpose cloning region and back into the βgalactosidase gene. The short sequence at the right-hand side is that of the primer used to initiate DNA synthesis across the
cloned insert. The numbered boxes correspond to the amino acids of the β-galactosidase fragment.
dideoxynucleotides as substrates (Fig. 7.1). Once the
analogue is incorporated at the growing point of the
DNA chain, the 3′ end lacks a hydroxyl group and
is no longer a substrate for chain elongation. Thus,
the growing DNA chain is terminated, i.e. dideoxynucleotides act as chain terminators. In practice,
the Klenow fragment of DNA polymerase is used
because this lacks the 5′ → 3′ exonuclease activity
associated with the intact enzyme. Initiation of DNA
synthesis requires a primer and this is usually a
chemically synthesized oligonucleotide which is
annealed close to the sequence being analysed.
POGC07 9/11/2001 11:07 AM Page 123
Sequencing and mutagenesis
DNA synthesis is carried out in the presence of the
four deoxynucleoside triphosphates, one or more of
which is labelled with 32P, and in four separate incubation mixes containing a low concentration of one
each of the four dideoxynucleoside triphosphate
analogues. Therefore, in each reaction there is a
population of partially synthesized radioactive DNA
molecules, each having a common 5′ end, but each
varying in length to a base-specific 3′ end (Fig. 7.2).
After a suitable incubation period, the DNA in each
mixture is denatured and electrophoresed in a
sequencing gel.
A sequencing gel is a high-resolution gel designed
to fractionate single-stranded (denatured) DNA fragments on the basis of their size and which is capable
of resolving fragments differing in length by a single
base pair. They routinely contain 6–20% polyacrylamide and 7 mol/l urea. The function of the urea is
to minimize DNA secondary structure, which affects
electrophoretic mobility. The gel is run at sufficient
power to heat up to about 70°C. This also minimizes
DNA secondary structure. The labelled DNA bands
obtained after such electrophoresis are revealed by
autoradiography on large sheets of X-ray film and
from these the sequence can be read (Fig. 7.3).
To facilitate the isolation of single strands, the DNA
to be sequenced may be cloned into one of the clustered cloning sites in the lac region of the M13 mp
series of vectors (Fig. 7.4). A feature of these vectors
is that cloning into the same region can be mediated
by any one of a large selection of restriction enzymes
but still permits the use of a single sequencing primer.
Modifications of chain-terminator
sequencing
The sharpness of the autoradiographic images can
be improved by replacing the 32P-radiolabel with the
much lower-energy 33P or 35S. In the case of 35S, this
is achieved by including an α-35S-deoxynucleoside
triphosphate (Fig. 7.5) in the sequencing reaction.
This modified nucleotide is accepted by DNA polymerase and incorporated into the growing DNA
chain. Non-isotopic detection methods have also been
developed with chemiluminescent, chromogenic or
fluorogenic reporter systems. Although the sensitivity of these methods is not as great as with radiolabels, it is adequate for many purposes.
γ
β
O
HO
P
O–
123
α
35
S
O
O
P
O–
O
P
O
CH2
O–
Base
O
H
H
H
H
OH
H
Fig. 7.5 Structure of an α-35S-deoxynucleoside triphosphate.
Other technical improvements to Sanger’s original
method have been made by replacing the Klenow
fragment of Escherichia coli DNA polymerase I.
Natural or modified forms of the phage T7 DNA
polymerase (‘Sequenase’) have found favour, as has
the DNA polymerase of the thermophilic bacterium
Thermus aquaticus (Taq DNA polymerase). The T7
DNA polymerase is more processive than Klenow
polymerase, i.e. it is capable of polymerizing a longer
run of nucleotides before releasing them from the
template. Also, its incorporation of dideoxynucleotides is less affected by local nucleotide sequences
and so the sequencing ladders comprise a series of
bands with more even intensities. The Taq DNA
polymerase can be used in a chain-termination reaction carried out at high temperatures (65–70°C) and
this minimizes chain-termination artefacts caused by
secondary structure in the DNA. Tabor and Richardson (1995) have shown that replacing a single
phenylalanine residue of Taq DNA polymerase with
a tyrosine residue results in a thermostable sequencing enzyme that no longer discriminates between
dideoxy- and deoxynucleotides.
The combination of chain-terminator sequencing
and M13 vectors to produce single-stranded DNA
is very powerful. Very good-quality sequencing is
obtainable with this technique, especially when
the improvements given by 35S-labelled precursors
and T7 DNA polymerase are exploited. Further
modifications allow sequencing of ‘double-stranded’
DNA, i.e. double-stranded input DNA is denatured
by alkali and neutralized and then one strand is
annealed with a specific primer for the actual chainterminator sequencing reactions. This approach has
gained in popularity as the convenience of having
a universal primer has grown less important with
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CHAPTER 7
the widespread availability of oligonucleotide synthesizers. With this development, Sanger sequencing has been liberated from its attachment to the M13
cloning system; for example, polymerase chain
reaction (PCR)-amplified DNA segments can be
sequenced directly. One variant of the doublestranded approach, often employed in automated
sequencing, is ‘cycle sequencing’. This involves a
linear amplification of the sequencing reaction,
using 25 cycles of denaturation, annealing of a
specific primer to one strand only and extension in
the presence of Taq DNA polymerase plus labelled
dideoxynucleotides. Alternatively, labelled primers
can be used with unlabelled dideoxynucleotides.
Automated DNA sequencing
In manual sequencing, the DNA fragments are
radiolabelled in four chain-termination reactions,
separated on the sequencing gel in four lanes and
detected by autoradiography. This approach is not
well suited to automation. To automate the process,
it is desirable to acquire sequence data in real time by
detecting the DNA bands within the gel during the
electrophoretic separation. However, this is not trivial, as there are only about 10−15−10−16 moles of
DNA per band. The solution to the detection problem is to use fluorescence methods. In practice, the
fluorescent tags are attached to the chain-terminating
nucleotides. Each of the four dideoxynucleotides
carries a spectrally different fluorophore. The tag is
incorporated into the DNA molecule by the DNA
polymerase and accomplishes two operations in one
step: it terminates synthesis and it attaches the
fluorophore to the end of the molecule. Alternatively, fluorescent primers can be used with nonlabelled dideoxynucleotides. By using four different
fluorescent dyes, it is possible to electrophorese
all four chain-terminating reactions together in
one lane of a sequencing gel. The DNA bands are
detected by their fluorescence as they electrophorese
past a detector (Fig. 7.6). If the detector is made to
V−
Upper buffer
reservoir
Polyacrylamide
sequencing
gel
Electrophoresis
Fluorescently
labelled
DNA bands
V+
Lower
buffer
reservoir
Scan line
Sequence
A G C T T A C GG A C T A A T G C
Dye 1
Dye 2
Dye 3
Dye 4
Computer
Time
Fluorescence
Fluorescence detector
Fig. 7.6 Block diagram of an
automated DNA sequencer and
idealized representation of the
correspondence between fluorescence
in a single electrophoresis lane and
nucleotide sequence.
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Sequencing and mutagenesis
scan horizontally across the base of a slab gel, many
separate sequences can be scanned, one sequence
per lane. Because the different fluorophores affect
the mobility of fragments to different extents, sophisticated software is incorporated into the scanning
step to ensure that bands are read in the correct
order. A simpler method is to use only one fluorophore and to run the different chain-terminating
reactions in different lanes.
For high-sensitivity DNA detection in four-colour
sequencing and high-accuracy base calling, one
would ideally like the following criteria to be met:
each of the four dyes to exhibit strong absorption at a
common laser wavelength; to have an emission
maximum at a distinctly different wavelength; and
to introduce the same relative mobility shift of the
DNA sequencing fragments. Recently, dyes with
these properties have been identified and successfully applied to automated sequencing (Glazer &
Mathies 1997).
Automated DNA sequencers offer a number of
advantages that are not particularly obvious. First,
manual sequencing can generate excellent data, but
even in the best sequencing laboratories poor
autoradiographs are frequently produced that make
sequence reading difficult or impossible. Usually the
problem is related to the need to run different termination reactions in different tracks of the gel. Skilled
DNA sequencers ignore bad sequencing tracks, but
many laboratories do not. This leads to poor-quality
sequence data. The use of a single-gel track for all
four dideoxy reactions means that this problem is
less acute in automated sequencing. Nevertheless, it
is desirable to sequence a piece of DNA several times
and on both strands, to eliminate errors caused by
technical problems. It should be noted that long
runs of the same nucleotide or a high G+C content
can cause compression of the bands on a gel, necessitating manual reading of the data, even with
an automated system. Note also that multiple,
tandem short repeats, which are common in the
DNA of higher eukaryotes, can reduce the fidelity
of DNA copying, particularly with Taq DNA polymerase. The second advantage of automated DNA
sequencers is that the output from them is in
machine-readable form. This eliminates the errors
that arise when DNA sequences are read and transcribed manually.
125
A third advantage derives from the new generation of sequencers that have been introduced
recently. In these sequencers, the slab gel is replaced
with 48 or 96 capillaries filled with the gel matrix.
The key feature of this system is that the equipment
has been designed for use with robotics, thereby
minimizing hands-on time and increasing throughput. With a 96-capillary sequencer, it is possible to
sequence up to 750 000 nucleotides per day.
Sequence accuracy
As part of a programme to sequence a 96 kb stretch
of mouse DNA, Wilson et al. (1992) analysed 288
sequences containing part of the vector DNA. By
comparing raw sequence data with known vector
sequences, it was possible to calculate the error
frequency. Sequences that were read beyond 400 bp
contained an average of 3.2% error, while those less
than 400 bp had 2.8% error. At least one-third of
the errors were due to ambiguities in sequence reading. In those sequences longer than 400 bp that
were read, most errors occurred late in the sequence
and were often present as extra bases in a run of
two or more of the same nucleotide. The remainder
of the errors were due to secondary structure in
the template DNA. However, because the complete
sequence was analysed with an average 5.9-fold
redundancy and most of it on both strands, the final
error frequency is estimated to be less than 0.1%. In
comparison, 35 different European laboratories were
engaged in sequencing the Saccharomyces cerevisiae
genome, with the attendant possibility of a very high
error frequency. However, by using a DNA coordinator who implemented quality-control procedures
(Table 7.1), the overall sequence accuracy for yeast
chromosome XI (666 448 bp) was estimated to
be 99.97% (Dujon et al. 1994), i.e. similar to that
noted above. Lipshutz et al. (1994) have described a
software program for estimating DNA sequence
confidence. Fabret et al. (1995) have analysed the
errors in finished sequences. They took advantage of
the fact that the surfactin operon of Bacillus subtilis
had been sequenced by three independent groups.
This enabled the actual error rate to be calculated. It
was found to range from 0.02 to 0.27%, the different
error rates being ascribed to the detailed sequencing tactics used. Other studies of DNA sequencing
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CHAPTER 7
Total number
of fragments
Total bp
verified
Error %
detected
Original overlap between cosmids
28
63 424
0.02
Resequencing of selected segments
(3–5 kb long)
21
72 270
0.03
Resequencing of random segments
(∼300 bp long)
71
18 778
0.05
Resequencing of suspected segments
(∼300 bp long) from designed
oligonucleotide pairs
60
17 035
0.03
180
171 507
Method of verification
Total
Average error rate
accuracy have been summarized by Yager et al.
(1997). In general, a sequence read of > 350 nucleotides at 99% accuracy can be expected using current
ultrathin-slab gel technology. Reading lengths in
excess of 1000 nucleotides have been reported
(Noolandi et al. 1993; Voss et al. 1995).
Whole-genome sequencing
As noted in Chapter 1 (p. 2), many different genomes
have been completely sequenced and the list includes
viruses, bacteria, yeast, Caenorhabditis, Drosophila,
Arabidopsis and humans. A detailed description of
the methodology used for sequencing these genomes
is outside the scope of this book and the interested
reader is referred to the sister publication Principles
of Genome Analysis (Primrose & Twyman 2002).
Suffice is to say that the underlying principle is to
subdivide the genome into small fragments of a
size suitable for sequencing by the methods just
described. Provided that the fragments overlap, the
individual sequences can then be assembled into
the complete genome sequence. However, the scale
of the task of complete sequence assembly can be
gauged from a comparison of the length of fragment that can be sequenced (600–1000 nucleotides)
with the number of such fragments in the genome
(> 3 million for the human genome)!
Table 7.1 Quality control of sequence
data from 35 laboratories engaged in
sequencing yeast chromosome XI.
(Data from Dujon et al. 1994 with
permission from Nature, © Macmillan
Magazines Ltd.)
0.03
Analysing sequence data
DNA sequence databases
Since the current DNA sequencing technology was
developed, a large amount of DNA sequence data
has accumulated. These data are maintained in
three databases: the National Center for Biotechnology Information in the USA, the DNA Databank
of Japan and the European Bioinformatic Institute
in the UK (Benson et al. 1996, 1997, Stoesser et al.
1997, Tateno & Gojobori 1997). Each of these
three groups collects a portion of the total sequence
data reported worldwide and all new and updated
database entries are exchanged between the groups
on a daily basis. In addition, several specialized
genome databases exist, including seven for bacterial genomes: four for E. coli, two for B. subtilis and
one at the Institute for Genome Research, an organization responsible for the complete sequencing of a
number of genomes. Users worldwide can access
these databases directly via the Worldwide Web or
receive the information on CD-ROMs. The former
option is the best because it ensures that an up-todate database is being used. There are a number of
different sequence-retrieval systems and the best of
these are Network Entrez and DNA Workbench
(Brenner 1995).
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Sequencing and mutagenesis
In working with databases, it is important to recognize their inherent deficiencies. As well as errors
in the sequencing process itself, there can be transcriptional errors when the data are transferred
from laboratory notebook to publications and databases. For example, when screening 300 human
protein sequences in the SWISS-PROT database that
had been published separately more than once, Bork
(1996) found that 0.3% of the amino acids were
different. This is a lower limit, for many corrections
will already have been made and in many instances
the sequences appearing in two different publications are not independent. Note that only stop
codons and frame shifts can be detected unambiguously: point mutations are hard to verify, as natural
polymorphisms or strain differences cannot easily be
excluded. Sequencing by hybridization may be of
great use here. Other database problems include
misspelling of genes, resulting in confusion with
ones of similar name or the generation of synonyms:
different genes being given the same name and
multiple synonyms for the same gene. Examples of
the latter are the E. coli gene hns, which has eight
synonyms, and the protein annexin V, which has
five synonyms.
Analysing sequence data
Discovering new genes and their functions is a key
step in analysing new sequence data. The process is
facilitated by special-purpose gene-finding software,
by searches in key databases and by programs for
finding particular sites relevant to gene expression,
e.g. splice sites and promoters. Unfortunately, no
one software package contains all the necessary
tools. Rather, optimal gene finding is dependent on
combining evidence derived from use of multiple
software tools (Table 7.2).
Fickett (1996) has described a framework for
finding genes which makes use of a number of different software programs. Evidence for the presence or
absence of genes in a sequence is gathered from a
number of sources. These include sequence similarity
to other features, such as repeats, which are unlikely
to overlap protein-coding sequences, sequence similarity to other genes, statistical regularity evincing
apparent codon bias over a region, and matches to
template patterns for functional sites on the DNA,
127
such as consensus sequences for TATA boxes. All
the information so gathered is integrated to make as
coherent a picture as possible.
When analysing sequences from eukaryotes, it is
best to locate and remove interspersed repeats before
searching for genes. Not only can repeats confuse
other analyses, such as database searches, but they
provide important negative information on the location of gene features. For example, such repeats
rarely overlap the promoters or coding portions of
exons. Once this is done, the next step is to identify
open reading frames (ORFs). Despite the availability
of sophisticated software search routines, unambiguous assignment of ORFs is not easy. For example, the Haemophilus influenzae genome sequence
submitted to the database included 1747 predicted
protein-encoding genes (Fleischmann et al. 1995).
When Tatusov et al. (1996) reanalysed the sequence
data using different algorithms and different discrimination criteria, they identified a new set of
1703 putative protein-encoding genes. In addition
to 1572 ORFs, which remained the same, they
identified 23 new ORFs, modified 107 others and
discarded the balance. Note that gene finding
is relatively easy in compact and almost intron-free
genomes, such as yeast. In higher plants and animals, the task is much greater, for a 2 kb ORF could
be split into 15 exons spread over 30 kb of genomic
DNA.
Database searches
Searching for a known homologue is the most
widely used means of identifying genes in a new
sequence. If a putative protein encoded by an
uncharacterized ORF shows statistically significant
similarity to another protein of known function, this
simultaneously proves, beyond doubt, that the ORF
in question is a bona fide new gene and predicts its
likely function. Even if the homologue of the new
protein has not been characterized, useful information is produced in the form of conserved motifs that
may be important for protein function. In this way,
Koonin et al. (1994) analysed the information contained in the complete sequence of yeast chromosome III and found that 61% of the probable gene
products had significant similarities to sequences in
the current databases. As many as 54% of them had
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CHAPTER 7
Table 7.2 Internet tools for gene discovery in DNA sequence data.
Important Home Pages
European Molecular Biology Laboratory (EMBL)
http://www.ebi.ac.uk/ebi_docs/embl_db/ebi/topembl.html
Cambridge, UK.
UK Human Genome Mapping Project – Resource Center (HGMP-RC)
http://www.hgmp.mrc.ac.uk/
SeqNet: UK Node of European Molecular Biology Network (EMBNet)
http://www.seqnet.dl.ac.uk/About/SEQNET/
GenBank
http://www.ncbi.nlm.nih.gov/Genbank/
GenBank at the National Center for Biotechnology (NCBI) of the National Library of Medicine (NLM) at the National Institutes for
Health (NIH) campus, USA.
DNA Databank of Japan (DDBJ)
http://www.ddbj.nig.ac.jp
Genome Sequence DataBase (GSDB)
http://seqsim.ncgr.org/
The National Center for Genome Resources, Genome Sequence Database.
The server is a supercomputer with genomic algorithm acceleration.
The Institute for Genomic Research (TIGR)
http://www.tigr.org/
The Sanger Centre
http://www.sanger.ac.uk/
Swiss Institute of Bioinformatics (Expasy)
http://www.expasy.ch/
Genomic Sequence Databases
These are the three primary sequence databases. They exchange sequence information daily.
European Molecular Biology Laboratory (EMBL)
http://www2.ebi.ac.uk/Help/General/general.html
Cambridge, UK.
GenBank
http://www.ncbi.nlm.nih.gov/Web/Search/index.html
GenBank at the National Center for Biotechnology (NCBI) of the National Library of Medicine (NLM) at the National Institutes for
Health (NIH) campus, USA.
DNA Databank of Japan (DDBJ)
http://www.ddbj.nig.ac.jp
Sequence Search and Retrieval
Sequence Retrieval System (SRS)
http://srs.hgmp.mrc.ac.uk/
Entrez
http://www.ncbi.nlm.nih.gov/Entrez/
Very comprehensive indeed!
Open Reading Frame (ORF) Finder
ORF Finder
http://www.ncbi.nlm.nih.gov/gorf/gorf.html
Finds likely open reading frames in a sequence.
continued
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Sequencing and mutagenesis
129
Table 7.2 (cont’d )
Sequence Translation
Nucleotide to Protein (EMBL)
5-4-1-DUW/UK
http://www.ebi.ac.uk/contrib/tommaso/translate.html
Translates a nucleotide sequence into a protein sequence.
Forward and Reverse Translation
5-5-2-DUPUFWE/UK
http://www.sanger.ac.uk/Software/Wise2/genewiseform.shtml
Translates a protein sequence into a genomic DNA sequence, and vice versa.
Protein and cDNA Translation
5-4-2-DcUPU/UK
http://www.sanger.ac.uk/Software/Wise2/genewiseform.shtml
Translates a protein sequence into a cDNA sequence, and vice versa.
Protein Sequence Databases
Translated EMBL (TrEMBL)
http://www.expasy.ch/sprot/sprot-top.html
Database of all the protein-coding regions stored in the EMBL database.
Comprehensive, but (generally) at the cost of poor annotation.
SWISS-PROT
http://expasy.hcuge.ch/sprot/sprot-top.html
A database of protein sequences, translated from the EMBL genomic database. Protein sequences have been checked and
annotated.
PIR
http://www nbrf.georgetown.edu/pir/
Four databases: PIR1 is the most comprehensive with entries classified and annotated. PIR4 is the least comprehensive, with
unencoded or untranslated entries.
Protein Three-dimensional Structure Database
Protein Databank
http://pdb.pdb.bnl.gov/
Three-dimensional protein structures which can be downloaded and viewed locally (viewer required) or viewed in a hypertext
browser window (e.g. Netscape). The structures are experimentally determined by X-ray crystallography and nuclear magnetic
resonance (NMR) imaging.
Protein Analysis Utilities
Web Cutter – Restriction Enzyme Mapping Utility
http://www.medkem.gu.se/cutter/
Map restriction-enzyme sites on your sequence. Easy to use and comprehensive options.
Sequence Identification (BLAST), FASTA, etc.)
The Sanger Centre Database Search Services
5-2-5-DDPDFWE/UK – Clean, simple design.
http://www.sanger.ac.uk/DataSearch/
BLAST and WU-BLAST 2.0 searches which can be refined to finished and/or unfinished genomic sequences.
BLAST 2 Similarity Search (EMBNet)
5-3-2-PUWE/Switzerland
http://www.ch.embnet.org/software/frameBLAST.html
WU-BLAST 2.0 similarity searches.
FASTA 3 (EMBL)
http://www2.ebi.ac.uk/fasta3/
FASTA 3 similarity search. Links to GenBank sequences returned. MolView presents alignment results well and in colour.
continued
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CHAPTER 7
Table 7.2 (cont’d )
FASTA
http://www2.igh.cnrs.fr/bin/fasta-guess.cgi
FASTA similarity search and a clean, basic and simple interface.
Blitz
http://www.ebi.ac.uk/searches/blitz.html
Blitz should only be used if BLAST and FASTA have failed to yield the desired results. It is a sensitive search, but also slow!
Beauty
http://dot.imgen.bcm.tmc.edu:9331/seq-search/protein-search.html
Beauty is an enhanced BLAST search, returning output which predicts the function of the protein being tested.
Sequence Alignment
Network Protein Sequence Analysis
5-4-1-PUW/France
http://pbil.ibcp.fr/NPSA/npsa_clustalw.html
ClustalW multiple sequence alignment.
Multiple Sequence Alignment with Hierarchical Clustering
5-5-3-PUW/France
http://www.toulouse.inra.fr/multalin.html
Sequence alignment with a colour output where differing or similar amino acids in the alignment can be highlighted.
ALIGN
5-1-1-DUPUWE/France
http://www2.igh.cnrs.fr/bin/align-guess.cgi
Applies the BLOSUM50 matrix to deduce the optimal alignment between two sequences.
New Sequence Submission
EMBL
http://www.ebi.ac.uk/Submissions/index.html
BankIt (GenBank)
http://www.ncbi.nlm.nih.gov/BankIt/
known functions or were related to functionally
characterized proteins and 19% were similar to proteins of known three-dimensional structure. Other
examples are given later. The methods of choice for
screening databases in this way are those such as
BLASTX that translate the query nucleotide sequence
in all six reading frames and compare the resulting
putative protein sequences to the protein-sequence
database. Such methods allow the detection of
frame-shift errors and will not miss even small ORFs
if homologues are present in the database.
Significant matches of a novel ORF to another
sequence may be in any of four classes (Oliver
1996). First, a match may predict both the biochemical and physiological function of the novel
gene. An example is ORF YCR24c from yeast chromosome III and the E. coli Asn-tRNA synthetase.
Secondly, a match may define the biochemical function of a gene product without revealing its biochemical function. An example of this is the five
protein-kinase genes found on yeast chromosome III
whose biochemical function is clear (they phosphorylate proteins) but whose particular physiological
function in yeast is unknown. Thirdly, a match may
be to a gene from another organism whose function
is unknown in that organism, e.g. ORF YCR63w from
yeast, protein G10 from Xenopus and novel genes
from Caenorhabditis and humans, none of whose
functions is known. Finally, a match may occur to a
gene of known function that merely reveals that our
understanding of that function is very superficial,
e.g. yeast ORF YCL17c and the Nif S protein of nitrogen-fixing bacteria. After similar sequences were
found in a number of bacteria which do not fix nitro-
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Sequencing and mutagenesis
1
Fraction of error - free genes
gen, it was shown that the Nif S protein is a pyridoxal
phosphate-dependent aminotransferase.
Because the DNA and protein-sequence databases
are updated daily, it could be that homologues of
previously unidentified proteins have been found.
Thus it could pay to repeat the search of the
databases at regular intervals, as Robinson et al.
(1994) found. Starting with more than 18 million
bp of prokaryotic DNA sequence, they executed a
systematic search for previously undetected proteincoding genes. They removed all DNA regions known
to encode proteins or structural RNAs and used an
algorithm to translate the remaining DNA in all six
reading frames. A search of the resultant translations against the protein-sequence databases uncovered more than 450 genes which previously had
escaped detection. Seven of these genes belonged
to gene families not previously identified in prokaryotes. Others belonged to gene families with critical
roles in metabolism. Clearly, periodic exhaustive
reappraisal of databases can be very productive!
If homology with known proteins is to be used to
analyse sequence data, then sequence accuracy is
paramount. With the gene density and ORF size
distribution of yeast (Dujon et al. 1994) and the
nematode (Wilson et al. 1994), even relatively rare
sequencing errors, many of which are missing
nucleotides, will result in a large fraction of the
protein-coding genes being affected by frame shifts.
At 99% accuracy virtually all genes will contain
errors and at 99.9% accuracy two-thirds of the
genes will still contain an error, most probably a
frame shift. With the 99.97% accuracy obtained in
the yeast sequencing, about one-third of predicted
genes will still contain errors (Fig. 7.7).
Because sequence-similarity search programs are
so vital to the analysis of DNA sequence data, much
attention has been paid to the precise algorithms
used and their precise speed. However, the effectiveness of these searches is dependent on a number of
factors. These include the choice of scoring system,
the statistical significance of alignments and the
nature and extent of sequence redundancy in the
databases (Altschul et al. 1994; Coulson 1994).
Most database search algorithms rank alignments
by a score whose calculation is dependent upon a
particular scoring system. Optimal strategies for
detecting similarities between DNA protein-coding
regions differ from those for non-coding regions.
131
2
0.5
1
0
10
1
0.1
0.01
0.001
Average sequence-error rate (%)
Fig. 7.7 Quantitative effect of sequence accuracy on gene
accuracy. The figure indicates the fraction of error-free genes
predicted from a DNA sequence, as a function of average
sequence accuracy. The theoretical curve was computed
using the size distribution of protein-coding genes in yeast
and assuming random occurrence of sequencing errors. All
types of errors are considered together. In practice, frame
shifts, which have the most deleterious effects on gene
interpretation, represent about two-thirds of all sequencing
errors. The figure illustrates the difference between an
average sequence accuracy of 99.9%, where only one-third
of all genes are properly described, and an average sequence
accuracy of 99.99%, where more than 85% of genes are
properly described. It also illustrates the difference between
average database entries (99.9% accuracy, arrow 1) and
systematic sequencing programs with sequence verifications
(99.97% accuracy, arrow 2). (Redrawn with permission from
Dujon 1996.)
Special scoring systems have been developed for
detecting frame-shift errors in the databases, a
problem highlighted above. Thus a database search
program should make use of a variety of scoring systems. Furthermore, given a query sequence, most
database search programs will produce an ordered
list of imperfectly matching database similarities,
but none of them need have any biological significance. Many of the genes originally predicted by
these statistical methods have subsequently proved
to be homologous to newly described genes or have
been confirmed experimentally, thus supporting the
robustness of the predictive methods (Koonin et al.
1996).
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CHAPTER 7
Eventually, with the accumulation of new
sequences, sequence conservation will become the
definitive criterion for gene identification and the
contribution of statistical methods will decrease.
Nevertheless, it is still likely that some genes will not
have identifiable homologues and statistical and experimental approaches will remain necessary for their
detection. Furthermore, even for genes that have
homologues, statistical methods of coding-potential
analysis will remain useful for localizing frame shifts
and choosing among the possible initiation codons.
Changing genes: site-directed
mutagenesis
Three different methods of site-directed mutagenesis
have been devised: cassette mutagenesis, primer
extension and procedures based on the PCR. All
three are described below but the reader wishing
more detail should consult the review of Ling and
Robinson (1998).
In some cases, the goal of protein engineering is to
generate a molecule with an improvement in some
operating parameter, but it is not known what
amino acid changes to make. In this situation,
a random mutagenesis strategy provides a route to
the desired protein. However, methods based on
gene manipulation differ from traditional mutagenesis in that the mutations are restricted to the gene
of interest or a small portion of it. Genetic engineering also provides a number of simple methods of
generating chimeric proteins where each domain
is derived from a different protein.
It should not be forgotten that constructing the
mutant DNA is only part of the task. The vector for
expression, the expression system and strategies
for purification and assay must also be considered
before embarking on protein mutagenesis.
Cassette mutagenesis
In cassette mutagenesis, a synthetic DNA fragment
containing the desired mutant sequence is used to
replace the corresponding sequence in the wild-type
gene. This method was originally used to generate
improved variants of the enzyme subtilisin (Wells
et al. 1985). It is a simple method for which the
efficiency of mutagenesis is close to 100%. The
disadvantages are the requirement for unique
restriction sites flanking the region of interest and
the limitation on the realistic number of different
oligonucleotide replacements that can be synthesized. The latter problem can be minimized by the use
of doped oligonucleotides (Fig. 7.8; Reidhaar-Olson
and Sauer, 1988), as described on pp. 107–109.
Primer extension:
the single-primer method
The simplest method of site-directed mutagenesis is
the single-primer method (Gillam et al. 1980, Zoller
& Smith 1983). The method involves priming in
vitro DNA synthesis with a chemically synthesized
oligonucleotide (7–20 nucleotides long) that carries
a base mismatch with the complementary sequence.
As shown in Fig. 7.9, the method requires that the
DNA to be mutated is available in single-stranded
form, and cloning the gene in M13-based vectors
makes this easy. However, DNA cloned in a plasmid
and obtained in duplex form can also be converted to
a partially single-stranded molecule that is suitable
(Dalbadie-McFarland et al. 1982).
Arg Glu Ile * * * Glu Met * * * Glu Ala Val Ser Met
G
T CGA GAA A TC NNG
C GAG A TG NN C GAA GCG GT T AGC A TG
CT T T AG I I I CTC T AC I I I CT T CGC CAA TC
AGC T
XhoI
GTAC
SphI
Fig. 7.8 Mutagenesis by means of
doped oligonucleotides. During
synthesis of the upper strand of the
oligonucleotide, a mixture of all four
nucleotides is used at the positions
indicated by the letter N. When the
lower strand is synthesized, inosine
(I) is inserted at the positions shown.
The double-stranded oligonucleotide
is inserted into the relevant position
of the vector.
POGC07 9/11/2001 11:07 AM Page 133
Sequencing and mutagenesis
T
A
Chemically synthesized
oligonucleotide
T
*
A G G C
C
C
GTA G
*
Anneal
A
C
GTA G
A
A
Single-stranded
M13 recombinant
133
TCAGGCT
(1) DNA polymerase + 4dNTPs
(2) T4 DNA ligase + ATP
A
A
*
A G G C
C
GTA G
*
T
T
C
Transform E. coli
Wild-type
A
A
T
A G G C
C
GTA G
*
T
The synthetic oligonucleotide primes DNA synthesis and is itself incorporated into the resulting
heteroduplex molecule. After transformation of the
host E. coli, this heteroduplex gives rise to homoduplexes whose sequences are either that of the original
wild-type DNA or that containing the mutated base.
The frequency with which mutated clones arise,
compared with wild-type clones, may be low. In
order to pick out mutants, the clones can be screened
by nucleic acid hybridization (see Chapter 6) with
32
P-labelled oligonucleotide as probe. Under suitable
conditions of stringency, i.e. temperature and cation
C
A
Fig. 7.9 Oligonucleotide-directed
mutagenesis. Asterisks indicate
mismatched bases. Originally the
Klenow fragment of DNA polymerase
was used, but now this has been largely
replaced with T7 polymerase.
A
A G G C
C
GTA G
*
T
T
C
Mutant
Screen plaques with 32P-labelled
oligonucleotide as hybridization
probe
Isolate mutant
concentration, a positive signal will be obtained only
with mutant clones. This allows ready detection of
the desired mutant (Wallace et al. 1981, Traboni
et al. 1983). In order to check that the procedure has
not introduced other adventitious changes, it is prudent to check the sequence of the mutant directly by
DNA sequencing. This was a particular necessity
with early versions of the technique which made use
of E. coli DNA polymerase. The more recent use of
the high-fidelity DNA polymerases from phages T4
and T7 has minimized the problem of extraneous
mutations, as well as shortening the time for copying
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CHAPTER 7
Multiple point
mutations
Insertion
mutagenesis
Deletion
mutagenesis
Mutant oligonucleotide
with multiple (four)
single base pair
mismatches
Mutant oligonucleotide
carrying a sequence to
be inserted sandwiched
between two regions
with sequences
complementary to sites
on either sides of the
target site in the
template
Mutant oligonucleotide
spanning the region to
be deleted, binding to
two separate sites, one
on either side of the
target
the second strand. Also, these polymerases do not
‘strand-displace’ the oligomer, a process which would
eliminate the original mutant oligonucleotide.
A variation of the procedure (Fig. 7.10) outlined
above involves oligonucleotides containing inserted
or deleted sequences. As long as stable hybrids are
formed with single-stranded wild-type DNA, priming of in vitro DNA synthesis can occur, ultimately
giving rise to clones corresponding to the inserted or
deleted sequence (Wallace et al. 1980, Norrander
et al. 1983).
Deficiencies of the single-primer method
The efficiency with which the single-primer method
yields mutants is dependent upon several factors.
The double-stranded heteroduplex molecules that
are generated will be contaminated both by any
single-stranded non-mutant template DNA that
has remained uncopied and by partially doublestranded molecules. The presence of these species
considerably reduces the proportion of mutant
progeny. They can be removed by sucrose gradient
centrifugation or by agarose gel electrophoresis, but
this is time-consuming and inconvenient.
Following transformation and in vivo DNA synthesis, segregation of the two strands of the heteroduplex molecule can occur, yielding a mixed
population of mutant and non-mutant progeny.
Mutant progeny have to be purified away from
parental molecules, and this process is complicated
by the cell’s mismatch repair system. In theory, the
mismatch repair system should yield equal numbers
Fig. 7.10 Oligonucleotide-directed
mutagenesis used for multiple point
mutation, insertion mutagenesis and
deletion mutagenesis.
of mutant and non-mutant progeny, but in practice
mutants are counterselected. The major reason for
this low yield of mutant progeny is that the methyldirected mismatch repair system of E. coli favours
the repair of non-methylated DNA. In the cell, newly
synthesized DNA strands that have not yet been
methylated are preferentially repaired at the position
of the mismatch, thereby eliminating a mutation. In
a similar way, the non-methylated in vitro-generated
mutant strand is repaired by the cell so that the
majority of progeny are wild type (Kramer, B. et al.
1984). The problems associated with the mismatch
repair system can be overcome by using host strains
carrying the mutL, mutS or mutH mutations, which
prevent the methyl-directed repair of mismatches.
A heteroduplex molecule with one mutant and
one non-mutant strand must inevitably give rise to
both mutant and non-mutant progeny upon replication. It would be desirable to suppress the growth of
non-mutants, and various strategies have been
developed with this in mind (Kramer, B. 1984, Carter
et al. 1985, Kunkel 1985, Sayers & Eckstein 1991).
Another disadvantage of all of the primer extension methods is that they require a single-stranded
template. In contrast, with PCR-based mutagenesis
(see below) the template can be single-stranded or
double-stranded, circular or linear. In comparison
with single-stranded DNAs, double-stranded DNAs
are much easier to prepare. Also, gene inserts are in
general more stable with double-stranded DNAs.
These facts account for the observation that most
commercial mutagenesis kits use double-stranded
templates.
POGC07 9/11/2001 11:07 AM Page 135
Sequencing and mutagenesis
135
Primer A
3‘
Primer A‘
5‘
5‘
PCR
3‘
PCR
3‘
5‘
Primer B
5‘
Primer C
3‘
PCR
PCR
3‘
5‘
5‘
3‘
Mix, denature and anneal
DNA polymerase
3‘
5‘
Fig. 7.11 Site-directed mutagenesis by
means of the PCR. The steps shown in
the top-left corner of the diagram show
the basic PCR method of mutagenesis.
The bottom half of the figure shows
how the mutation can be moved to the
middle of a DNA molecule. Primers are
shown in bold and primers A and A′ are
complementary.
5‘
+
5‘
3‘
3‘
3‘
5‘
DNA polymerase
5‘
PCR methods of site-directed
mutagenesis
Early work on the development of the PCR method of
DNA amplification showed its potential for mutagenesis (Scharf et al. 1986). Single bases mismatched
between the amplification primer and the template
become incorporated into the template sequence
as a result of amplification (Fig. 7.11). Higuchi et al.
(1988) have described a variation of the basic method
which enables a mutation in a PCR-produced DNA
fragment to be introduced anywhere along its length.
Two primary PCR reactions produce two overlapping DNA fragments, both bearing the same mutation in the overlap region. The overlap in sequence
allows the fragments to hybridize (Fig. 7.11). One
of the two possible hybrids is extended by DNA
3‘
+
PCR with primers B and C
polymerase to produce a duplex fragment. The other
hybrid has recessed 5′ ends and, since it is not a
substrate for the polymerase, is effectively lost from
the reaction mixture. As with conventional primerextension mutagenesis, deletions and insertions can
also be created.
The method of Higuchi et al. (1988) requires four
primers and three PCRs (a pair of PCRs to amplify the
overlapping segments and a third PCR to fuse the
two segments). Sarkar and Sommer (1990) have
described a simpler method, which utilizes three
oligonucleotide primers to perform two rounds of
PCR. In this method, the product of the first PCR is
used as a megaprimer for the second PCR (Fig. 7.12).
The advantage of a PCR-based mutagenic protocol is that the desired mutation is obtained with
100% efficiency. There are two disadvantages. First,
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CHAPTER 7
PCR
Multiple rounds
of PCR
PCR
3‘
5‘
Megaprimer
5‘
3‘
PCR
3‘
5‘
the PCR product usually needs to be ligated into a
vector, although Sarkar and Sommer (1990) have
generated the mutant protein directly, using coupled in vitro transcription and translation. Secondly,
Taq polymerase copies DNA with low fidelity (see
p. 23). Thus the sequence of the entire amplified
segment generated by PCR mutagenesis must be
determined to ensure that there are no extraneous
mutations. Alternatively, thermostable polymerases
with improved fidelity can be used (Cariello et al.
1991, Lundberg et al. 1991, Mattila et al. 1991).
Selection of mutant peptides by phage
and phasmid display
In phage display, a segment of foreign DNA is
inserted into either a phagemid or an infectious
filamentous phage genome and expressed as a
fusion product with a phage coat protein. It is a
very powerful technique for selecting and engineering polypeptides with novel functions. The technique was developed first for the E. coli phage
M13 (Parmley & Smith 1988), but has since been
extended to other phages such as T4 and λ (Ren &
Black 1998, Santini et al. 1998).
The M13 phage particle consists of a single-stranded
DNA molecule surrounded by a coat consisting of
several thousand copies of the major coat protein,
5‘
3‘
Fig. 7.12 The megaprimer method of
mutagenesis. The mutant molecule
produced in the early rounds of PCR
acts as a primer (‘megaprimer’) for
later rounds of PCR.
P8. At one end of the particle are five copies each of
the two minor coat proteins P9 and P7 and at the
other end five copies each of P3 and P6. In early
examples of phage display, a random DNA cassette
(see above) was inserted into either the P3 or the P8
gene at the junction between the signal sequence
and the native peptide. E. coli transfected with the
recombinant DNA molecules secreted phage particles that displayed on their surface the amino acids
encoded by the foreign DNA. Particular phage
displaying peptide motifs with, for example, antibodybinding properties were isolated by affinity chromatography (Fig. 7.13). Several rounds of affinity
chromatography and phage propagation can be
used to further enrich for phage with the desired
binding characteristics. In this way, millions of random peptides have been screened for their ability to
bind to an anti-peptide antibody or to streptavidin
(Cwirla et al. 1990, Devlin et al. 1990, Scott & Smith
1990), and variants of human growth hormone
with improved affinity and receptor specificity have
been isolated (Lowman et al. 1991).
One disadvantage of the original method of phage
display is that polypeptide inserts greater than 10
residues compromise coat-protein function and so
cannot be efficiently displayed. This problem can be
solved by the use of phagemid display (Bass et al.
1990). In this system, the starting-point is a plasmid
POGC07 9/11/2001 11:07 AM Page 137
Sequencing and mutagenesis
Promoter
Signal
137
Random
peptide
DNA
M13 gene III
Phage
vector
f1ori
Infect male E. coli
Phage
particle
Gene III
protein
DNA
Fig. 7.13 The principle of phage display
of random peptides.
carrying a single copy of the P3 or P8 gene from
M13 plus the M13 ori sequence (i.e. a phagemid,
see p. 70). As before, the random DNA sequence is
inserted into the P3 or P8 gene downstream from
the signal peptide-cleavage site and the construct
transformed into E. coli. Phage particles displaying
the amino acid sequences encoded by the DNA insert
are obtained by superinfecting the transformed cells
with helper phage. The resulting phage particles are
phenotypically mixed and their surfaces are a
mosaic of normal coat protein and fusion protein.
Specialized phagemid display vectors have been
developed for particular purposes. For example,
phagemids have been constructed that have an
amber (chain-terminating) codon immediately downstream from the foreign DNA insert and upstream
Random
protein
Test binding to antibody
or receptor
from the body of P3 or P8. When the recombinant
phagemid is transformed into non-suppressing
strains of E. coli, the protein encoded by the foreign
DNA terminates at the amber codon and is secreted
into the medium. However, if the phagemid is
transformed into cells carrying an amber suppressor, the entire fusion protein is synthesized and
displayed on the surface of the secreted phage particles (Winter et al. 1994). Other studies (Jespers
et al. 1995, Fuh & Sidhu 2000, Fuh et al. 2000) have
shown that proteins can be displayed as fusions to
the carboxy terminus of P3, P6 and P8. Although
amino-terminal display formats are likely to dominate established applications, carboxy-terminal display
permits constructs that are unsuited to aminoterminal display.
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CHAPTER 7
For a detailed review of phage and phagemid display, the reader should consult Sidhu (2000) and
Sidhu et al. (2000).
Directed mutation in vivo
In vitro mutagenesis methods are very useful when
working with small plasmids carrying cloned genes.
However, if it is desired to modify a single residue on
a very large vector, such as a yeast artificial chromosome (YAC), P1-derived artificial chromosome
(PAC) or bacterial artificial chromosome (BAC),
in vitro methods are not appropriate. With such
large plasmids, directed mutagenesis has to be done
in vivo and a number of different methods have
been described (Muyrers et al. 2000, Lalioti &
Heath 2001). The simplest is that of Yu et al. (2000),
which involves recombination between linear DNA
fragments carrying the desired mutation and the
homologous region on the chromosome.
Unlike many organisms, E. coli is not readily
transformed by linear DNA fragments. The main
reason for this is the rapid degradation of the DNA by
the intracellular RecBCD exonuclease. Mutant
strains lacking the exonuclease do not degrade linear DNA very rapidly, but such strains grow poorly,
are defective for recombination and do not support
efficient replication of plasmid vectors. The solution
devised by Yu et al. (2000) was to use an E. coli strain
containing a λ prophage harbouring the genes exo,
bet and gam under the control of a temperaturesensitive λcI repressor. The Gam gene product stops
the RecBCD nuclease from attacking the linear
DNA and the Exo and Bet gene products generate
recombination activity to enable DNA exchange
to occur.
POGC08 9/11/2001 11:06 AM Page 139
CH A P T E R 8
Cloning in bacteria other than
Escherichia coli
Introduction
Introducing DNA into bacterial cells
For many experiments it is convenient to use E. coli
as a recipient for genes cloned from eukaryotes or
other prokaryotes. Transformation is easy and there
is available a wide range of easy-to-use vectors
with specialist properties, e.g. regulatable high-level
gene expression. However, use of E. coli is not always
practicable because it lacks some auxiliary biochemical pathways that are essential for the phenotypic
expression of certain functions, e.g. degradation of
aromatic compounds, antibiotic synthesis, pathogenicity, sporulation, etc. In such circumstances,
the genes have to be cloned back into species similar
to those whence they were derived.
There are three prerequisites for cloning genes
in a new host. First, there needs to be a method for
introducing the DNA of interest into the potential
recipient. The methods available include transformation, conjugation and electroporation, and these
will be discussed in more detail later. Secondly, the
introduced DNA needs to be maintained in the
new host. Either it must function as a replicon in its
new environment or it has to be integrated into the
chromosome or a pre-existing plasmid. Finally, the
uptake and maintenance of the cloned genes will
only be detected if they are expressed. Thus the
inability to detect a cloned gene in a new bacterial
host could be due to failure to introduce the gene,
to maintain it or to express it, or to a combination
of these factors. Another cause of failure could
be restriction. For example, the frequency of transformation of Pseudomonas putida with plasmid
RSF1010 is 105 transformants/µg DNA, but only
if the plasmid is prepared from another P. putida
strain. Otherwise, no transformants are obtained
(Bagdasarian et al. 1 979). Wilkins et al. (1996) have
noted that conjugative transfer of promiscuous IncP
plasmids is unusually sensitive to restriction.
DNA can be transferred between different strains
of E. coli by the three classical methods of conjugation, transduction and transformation, as well as
by the newer method of electroporation. For genemanipulation work, transformation is nearly always
used. The reasons for this are threefold. First, it is relatively simple to do, particularly now that competent
cells are commercially available. Secondly, it can be
very efficient. Efficiencies of 108–109 transformants/
µg plasmid DNA are readily achievable and are more
than adequate for most applications. Thirdly, selftransmissible cloning vectors are much larger than
their non-transmissible counterparts because they
have to carry all the genes required for conjugal
transfer.
A large number of bacteria from different taxonomic groups, including archaebacteria, are known
to be transformable. For example, Lorenz and
Wackernagel (1994) listed over 40 species and the
numbers are still growing. However, transformation
in these organisms differs in a number of respects
from that in E. coli. First, transformation in these
organisms occurs naturally, whereas transformation
in E. coli is artificially induced. With the exception
of Neisseria gonorrhoeae, competence for transformation is a transient phenomenon. Secondly,
transformation can be sequence-independent, as in
Bacillus subtilis and Acinetobacter calcoaceticus, but in
other species (Haemophilus influenzae, N. gonorrhoeae)
is dependent on the presence of specific uptake
sequences. Thirdly, the mechanism of natural transformation involves breakage of the DNA duplex
and degradation of one of the two strands so that a
linear single strand can enter the cell (for review, see
Dubnau 1999). This mechanism is not compatible
with efficient plasmid transformation (see Box 8.1).
Geneticists working with B. subtilis and Streptomyces
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CHAPTER 8
Box 8.1 Transforming Bacillus subtilis with plasmid DNA
Although it is very easy to transform B. subtilis with
fragments of chromosomal DNA, there are problems
associated with transformation by plasmid molecules.
Ehrlich (1977) first reported that competent cultures
of B. subtilis can be transformed with covalently closed
circular (CCC) plasmid DNA from Staphylococcus aureus
and that this plasmid DNA is capable of autonomous
replication and expression in its new host. The
development of competence for transformation by
plasmid and chromosomal DNA follows a similar time
course and in both cases transformation is first-order
with respect to DNA concentration, suggesting that
a single DNA molecule is sufficient for successful
transformation (Contente & Dubnau 1979). However,
transformation of B. subtilis with plasmid DNA is
very inefficient in comparison with chromosomal
transformation, for only one transformant is obtained
per 103–104 plasmid molecules.
An explanation for the poor transformability of
plasmid DNA molecules was provided by Canosi et al.
(1978). They found that the specific activity of plasmid
DNA in transformation of B. subtilis was dependent on
the degree of oligomerization of the plasmid genome.
Purified monomeric CCC forms of plasmids transform
B. subtilis several orders of magnitude less efficiently
than do unfractionated plasmid preparations or
multimers. Furthermore, the low residual transforming
activity of monomeric CCC DNA molecules can be
attributed to low-level contamination with multimers
(Mottes et al. 1979). Using a recombinant plasmid
capable of replication in both E. coli and B. subtilis
(pHV14) (see p. 149), Mottes et al. (1979) were
able to show that plasmid transformation of E. coli
occurs regardless of the degree of oligomerization,
in contrast to the situation with B. subtilis.
Oligomerization of linearized plasmid DNA by DNA
ligase resulted in a substantial increase of specific
transforming activity when assayed with B. subtilis
and caused a decrease when used to transform
E. coli. An explanation of the molecular events in
transformation which generate the requirement for
oligomers has been presented by De Vos et al.
(1981). Basically, the plasmids are cleaved into linear
molecules upon contact with competent cells, just as
chromosomal DNA is cleaved during transformation of
Bacillus. Once the linear single-stranded form of the
plasmid enters the cell, it is not reproduced unless it
can circularize; hence the need for multimers to provide
regions of homology that can recombine. Michel et al.
(1982) have shown that multimers, or even dimers,
are not required, provided that part of the plasmid
genome is duplicated. They constructed plasmids
carrying direct internal repeats 260–2000 bp long
and found that circular or linear monomers of such
plasmids were active in transformation.
Canosi et al. (1981) have shown that plasmid
monomers will transform recombination-proficient
B. subtilis if they contain an insert of B. subtilis DNA.
However, the transformation efficiency of such
monomers is still considerably less than that of
oligomers. One consequence of the requirement for
plasmid oligomers for efficient transformation of
B. subtilis is that there have been very few successes
in obtaining large numbers of clones in B. subtilis
recipients (Keggins et al. 1978, Michel et al. 1980).
The potential for generating multimers during
ligation of vector and foreign DNA is limited.
Transformation by plasmid rescue
An alternative strategy for transforming B. subtilis has
been suggested by Gryczan et al. (1980). If plasmid
DNA is linearized by restriction-endonuclease cleavage,
no transformation of B. subtilis results. However, if
the recipient carries a homologous plasmid and if the
restriction cut occurs within a homologous marker,
then this same marker transforms efficiently. Since
this rescue of donor plasmid markers by a homologous
resident plasmid requires the B. subtilis recE gene
product, it must be due to recombination between
the linear donor DNA and the resident plasmid.
Since DNA linearized by restriction-endonuclease
cleavage at a unique site is monomeric, this rescue
system (plasmid rescue) bypasses the requirement
for a multimeric vector. The model presented by
De Vos et al. (1981) to explain the requirement for
oligomers (see above) can be adapted to account for
transformation by monomers by means of plasmid
continued
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Cloning in bacteria other than E. coli
141
Box 8.1 continued
rescue. In practice, foreign DNA is ligated to monomeric
vector DNA and the in vitro recombinants are used to
transform B. subtilis cells carrying a homologous
plasmid. Using such a ‘plasmid-rescue’ system,
Gryczan et al. (1980) were able to clone various
genes from B. licheniformis in B. subtilis.
One disadvantage of the plasmid-rescue method
is that transformants contain both the recombinant
molecule and the resident plasmid. Incompatibility
will result in segregation of the two plasmids. This
may require several subculture steps, although
Haima et al. (1990) observed very rapid segregation.
Alternatively, the recombinant plasmids can be
transformed into plasmid-free cells.
Transformation of protoplasts
A third method for plasmid DNA transformation in
B. subtilis involves polyethylene glycol (PEG) induction
of DNA uptake in protoplasts and subsequent
regeneration of the bacterial cell wall (Chang & Cohen
1979). The procedure is highly efficient and yields up
to 80% transformants, making the method suitable
for the introduction even of cryptic plasmids. In
addition to its much higher yield of plasmid-containing
transformants, the protoplast transformation system
differs in two respects from the ‘traditional’ system
using physiologically competent cells. First, linear
plasmid DNA and non-supercoiled circular plasmid
DNA molecules constructed by ligation in vitro can
be introduced at high efficiency into B. subtilis by
the protoplast transformation system, albeit at a
frequency 10–1000 lower than the frequency
observed for CCC plasmid DNA. However, the
efficiency of shotgun cloning is much lower with
protoplasts than with competent cells (Haima et al.
1988). Secondly, while competent cells can be
transformed easily for genetic determinants located
on the B. subtilis chromosome, no detectable
transformation with chromosomal DNA is seen using
the protoplast assay. Until recently, a disadvantage
of the protoplast system was that the regeneration
medium was nutritionally complex, necessitating
a two-step selection procedure for auxotrophic
markers. Details have been presented of a defined
regeneration medium by Puyet et al. 1987.
Table B8.1 Comparison of the different methods of transforming B. subtilis.
System
Efficiency
(transformants/mg DNA)
Advantages
Disadvantages
Competent
cells
Unfractionated plasmid
Linear
CCC monomer
CCC dimer
CCC multimer
2 × 104
0
4 × 104
8 × 103
2.6 × 105
Competent cells readily prepared
Transformants can be selected
readily on any medium
Recipient can be Rec −
Requires plasmid oligomers or
internally duplicated plasmids,
which makes shotgun experiments
difficult unless high DNA
concentrations and high vector/
donor DNA ratios are used
Not possible to use phosphatasetreated vector
Plasmid
rescue
Unfractionated plasmid
2 × 106
Oligomers not required
Can transform with linear DNA
Transformants can be selected on
any medium
Transformants contain resident
plasmid and incoming plasmid and
these have to be separated by
segregation or retransformation
Recipient must be Rec+
Protoplasts
Unfractionated plasmid
Linear
CCC monomer
CCC dimer
CCC multimer
3.8 × 106
2 × 104
3 × 106
2 × 106
2 × 106
Most efficient system
Gives up to 80% transformants
Does not require competent cells
Can transform with linear DNA and
can use phosphatase-treated vector
Efficiency lower with molecules
which have been cut and religated
Efficiency also very size-dependent,
and declines steeply as size increases
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CHAPTER 8
sp. have developed specialized methods for overcoming these problems. For work with other species,
electroporation (see p. 18) offers a much simpler
alternative, although the efficiency may not be
high e.g. in Methanococcus voltae, it was only 102
transformants/µg plasmid DNA (Tumbula & Whitman 1999).
Given that plasmid transformation is difficult in
many non-enteric bacteria, conjugation represents
an acceptable alternative. The term promiscuous
plasmids was originally coined for those plasmids
which are self-transmissible to a wide range of
other Gram-negative bacteria (see p. 45), where
they are stably maintained. The best examples are
the IncP alpha plasmids RP4. RP1, RK2, etc., which
are about 60 kb in size, but the IncW plasmid Sa
(29.6 kb) has been used extensively in Agrobacterium
tumefaciens. Self-transmissible plasmids carry tra
(transfer) genes encoding the conjugative apparatus.
IncP plasmids are able to promote the transfer of
other compatible plasmids, as well as themselves.
For example, they can mobilize IncQ plasmids,
such as RSF1010. More important, transfer can be
mediated between E. coli and Gram-positive bacteria
(Trieu-Cuot et al. 1987, Gormley & Davies 1991), as
well as to yeasts and fungi (Heinemann & Sprague
1989, Hayman & Bolen 1993, Bates et al. 1998).
However, the transfer range of a plasmid may be
greater than its replication maintenance or host
range (Mazodier & Davies 1991).
Self-transmissible plasmids have been identified in
many different Gram-positive genera. The transfer
regions of these plasmids are much smaller than
those for plasmids from Gram-negative bacteria. One
reason for this difference may be the much simpler
cell-wall structure in Gram-positive bacteria. As
well as being self-transmissible, many of these Grampositive plasmids are promiscuous. For example,
plasmid pAMβ1 was originally isolated from Enterococcus faecalis but can transfer to staphylococci,
streptococci, bacilli and lactic acid bacteria (De Vos
et al. 1997), as well as to Gram-negative bacteria,
such as E. coli (Trieu-Cuot et al. 1988). The selftransmissible plasmids of Gram-positive bacteria, like
their counterparts in the Gram-negative bacteria,
can also mobilize other plasmids between different
genera (Projan & Archer 1989, Charpentier et al.
1999). Non-self-transmissible plasmids can also be
mobilized within and between genera by conjugative transposons (Salyers et al. 1995, Charpentier
et al. 1999).
It should be noted that conjugation is not a
replacement for transformation or electroporation.
If DNA is manipulated in vitro, then it has to be
transferred into a host cell at some stage. In many
cases, this will be E. coli and transformation is a
suitable procedure. Once in E. coli, or any other
organism for that matter, it may be moved to other
bacteria directly by conjugation, as an alternative to
purifying the DNA and moving it by transformation
or electroporation.
Maintenance of recombinant
DNA in new hosts
For recombinant DNA to be maintained in a
new host cell, either it must be capable of replication or it must integrate into the chromosome or a
plasmid. In most instances, the recombinant will
be introduced as a covalently closed circle (CCC)
plasmid and maintenance will depend on the host
range of the plasmid. As noted in Chapter 4 (p. 45),
the host range of a plasmid is dependent on the
number of proteins required for its replication which
it encodes. Some plasmids have a very narrow host
range, whereas others can be maintained in a wide
range of Gram-negative or Gram-positive genera.
Some, such as the plasmids from Staphylococcus
aureus and RSF1010, can replicate in both Gramnegative and Gram-positive species (Lacks et al.
1986, Gormley & Davies 1991, Leenhouts et al.
1991).
As noted in Chapter 5, there is a very wide range
of specialist vectors for use in E. coli. However, most
of these vectors have a very narrow host range and
can be maintained only in enteric bacteria. A common way of extending the host range of these vectors is to form hybrids with plasmids from the target
species. The first such shuttle vectors to be described
were fusions between the E. coli vector pBR322 and
the S. aureus/B. subtilis plasmids pC194 and pUB110
(Ehrlich 1978). The advantage of shuttle plasmids is
that E. coli can be used as an efficient intermediate
host for cloning. This is particularly important if
transformation or electroporation of the alternative
host is very inefficient.
POGC08 9/11/2001 11:06 AM Page 143
Cloning in bacteria other than E. coli
Integration of recombinant DNA
When a recombinant plasmid is transferred to an
unrelated host cell, there are a number of possible
outcomes:
• It may be stably maintained as a plasmid.
• It may be lost.
• It may integrate into another replicon, usually the
chromosome.
• A gene on the plasmid may recombine with a
homologous gene elsewhere in the cell.
Under normal circumstances, a plasmid will be
maintained if it can replicate in the new host and will
be lost if it cannot. Plasmids which will be lost in
their new host are particularly useful for delivering
transposons (Saint et al. 1995, Maguin et al. 1996).
If the plasmid carries a cloned insert with homology
143
to a region of the chromosome, then the outcome
is quite different. The homologous region may be
excised and incorporated into the chromosome by
the normal recombination process, i.e. substitution
occurs via a double crossover event. If only a single
crossover occurs, the entire recombinant plasmid is
integrated into the chromosome.
It is possible to favour integration by transferring
a plasmid into a host in which it cannot replicate and
selecting for a plasmid-borne marker. For example,
Stoss et al. (1997) have constructed an integrative
vector by cloning a neomycin resistance gene and
part of the amyE (alpha amylase) gene of B. subtilis in
plasmid pBR322. When this plasmid is transferred
to B. subtilis, it is unable to replicate, but, if selection
is made for neomycin resistance, then integration
of the plasmid occurs at the amyE locus (Fig. 8.1).
ApR
Bam Bam
Bg/II
Bg/II
CmR
Bg/II
Bg/II
Recombination
BamHI
BamHI
CmR
Bg/II
ApR
Bg/II
Bg/II
Removal by Bg/II digestion
followed by ligation
Bg/II
BamHI
BamHI
Fig. 8.1 Cloning DNA sequences
flanking the site of insertion. The red
bar on the plasmid represents a BamHI
fragment of B. subtilis chromosomal
DNA carrying the amyE gene. Note that
the plasmid has no BglII sites. (See text
for details.)
CmR
ApR
POGC08 9/11/2001 11:06 AM Page 144
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CHAPTER 8
without tandem duplication of the chromosomal
segments. In this case, the plasmid DNA is linearized
before transformation, as shown in Fig. 8.2. The
same technique can be used to generate deletions.
The gene of interest is cloned, a portion of the gene
replaced in vitro with a fragment bearing an antibiotic marker and the linearized plasmid transformed into B. subtilis, with selection made for
antibiotic resistance.
l
Km
Z∆
ac
R
P
M15
Bg/II
B
C
Bg/II
A D
PstI
Km
∆M
15 P
Cut with PstI and transform
with selection for KmR
R
Z
lac
PstI
A
B
C
D
A
B
C
D
Bg/II
PstI
Plasmid
Chromosome
Fig. 8.2 Insertion of plasmid DNA into the chromosome by
a double crossover event. The B. subtilis DNA is shown in
grey and the letters A to D represent different chromosomal
sequences. Vector DNA is shown in white and other
vector-borne genes in pink.
This technique is particularly useful if one wishes
to construct a recombinant carrying a single copy of
a foreign gene.
Once a recombinant plasmid has integrated into
the chromosome, it is relatively easy to clone adjacent sequences. Suppose, for example, that a vector
carrying B. subtilis DNA in the BamHI site (Fig. 8.1)
has recombined into the chromosome. If the recombinant plasmid has no BglII sites, it can be recovered
by digesting the chromosomal DNA with BglII, ligating the resulting fragments and transforming E. coli
to ApR. However, the plasmid which is isolated will
be larger than the original one, because DNA flanking the site of insertion will also have been cloned.
In this way, Niaudet et al. (1982) used a plasmid
carrying a portion of the B. subtilis ilvA gene to clone
the adjacent thyA gene.
Genes cloned into a plasmid and flanked by regions
homologous to the chromosome can also integrate
Cloning in Gram-negative bacteria
other than E. coli
To clone DNA in non-enteric bacteria, a plasmid
cloning vehicle is required which can replicate in the
selected organism(s). Under normal circumstances,
E. coli will be used as an intermediate host for transformation of the ligation mix and screening for
recombinant plasmids. Therefore, the vector must
be able to replicate in E. coli as well. The options
which are available are to generate a shuttle vector
or to use a broad-host-range plasmid as a vector. If
a small plasmid can be isolated from the bacterium
of interest, then it is easy to splice it into an existing
E. coli vector to generate a shuttle vector. Recent
examples of this approach are the construction
of vectors for use in Pasteurella (Bills et al. 1993),
Desulfovibrio (Rousset et al. 1998) and Thermus (De
Grado et al. 1999). This approach is particularly
useful if the selectable markers used in E. coli
also function in the new host. Then one can take
advantage of the many different specialist vectors
(see Chapter 5) which already exist, e.g. expression vectors, secretion vectors, etc. If the selectable
markers are not expressed in the new host, then
extensive manipulations may be necessary just to
enable transformants to be detected.
With broad-host-range plasmids, there is a high
probability that the selectable markers will be expressed in the new host and confirming that this is
indeed the case is easy to do. However, the naturally
occurring plasmids do not fulfil all the criteria for an
ideal vector, which are:
• small size;
• having multiple selectable markers;
• having unique sites for a large number of restriction enzymes, preferably in genes with readily
scorable phenotypes.
POGC08 9/11/2001 11:06 AM Page 145
Cloning in bacteria other than E. coli
Consequently the natural plasmids have been extensively modified, but few approach the degree of
sophistication found in the standard E. coli vectors.
Vectors derived from the IncQ-group
plasmid RSF1010
Plasmid RSF1010 is a multicopy replicon which
specifies resistance to two antimicrobial agents,
sulphonamide and streptomycin. The plasmid DNA,
which is 8684 bp long, has been completely
sequenced (Scholz et al. 1989). A detailed physical
and functional map has been constructed (Bagdasarian et al. 1981, Scherzinger et al. 1984). The
features mapped are the restriction-endonuclease
recognition sites, RNA polymerase binding sites,
resistance determinants, genes for plasmid mobilization (mob), three replication proteins (Rep A, B and
C) and the origins of vegetative (ori) and transfer
(nic) replication.
Plasmid RSF1010 has unique cleavage sites for
EcoRI, BstEII, HpaI, DraII, NsiI and SacI and, from
the nucleotide sequence data, is predicted to have
unique sites for AfIIII, BanII, NotI, SacII, SfiI and
SplI. There are two PstI sites, about 750 bp apart,
which flank the sulphonamide-resistance determinant (Fig. 8.3). None of the unique cleavage sites is
HpaI
PstI
EcoRI
PstI
BstEII
SmR
SuR
0
8
7
A
B
1
2
RSF 1010
(8.68 kb)
6
3
5
C
145
located within the antibiotic-resistance determinants and none is particularly useful for cloning.
Before the Bst, Eco and Pst sites can be used, another
selective marker must be introduced into the
RSF1010 genome. This need arises because the SmR
and SuR genes are transcribed from the same promoter (Bagdasarian et al. 1981). Insertion of a DNA
fragment between the Pst sites inactivates both
resistance determinants. Although the Eco and Bst
sites lie outside the coding regions of the SmR gene,
streptomycin resistance is lost if a DNA fragment is
inserted at these sites unless the fragment provides a
new promoter. Furthermore, the SuR determinant
which remains is a poor selective marker.
A whole series of improved vectors has been
derived from RSF1010 but only a few are mentioned
here. The earliest vectors contained additional
unique cleavage sites and more useful antibioticresistance determinants. For example, plasmids
KT230 and KT231 encode KmR and SmR and have
unique sites for HindIII, XmaI, XhoRI and SstI which
can be used for insertional inactivation. These two
vectors have been used to clone in P. putida genes
involved in the catabolism of aromatic compounds
(Franklin et al. 1981). Vectors for the regulated
expression of cloned genes have also been described.
Some of these make use of the tac promoter
(Bagdasarian et al. 1983, Deretic et al. 1987) or the
phage T7 promoter (Davison et al. 1989), which will
function in P. putida as well as E. coli. Another makes
use of positively activated twin promoters from a
plasmid specifying catabolism of toluene and xylenes
(Mermod et al. 1986). Expression of cloned genes
can be obtained in a wide range of Gram-negative
bacteria following induction with micromolar quantities of benzoate, and the product of the cloned gene
can account for 5% of total cell protein.
ori
4
mob
Fig. 8.3 The structure of plasmid RSFI010. The pink tinted
areas show the positions of the SmR and SuR genes. The region
marked ori indicates the location of the origin of replication.
The mob function is required for conjugal mobilization by a
compatible self-transmissible plasmid. A, B and C are the
regions encoding the three replication proteins.
Vectors derived from the
IncP-group plasmids
Both the IncP alpha plasmids (R18, R68, RK2, RP1
and RP4), which are 60 kb in size, and the smaller
(52 kb) IncP beta plasmid R751 have been completely sequenced (Pansegrau et al. 1994, Thorsted
et al. 1998). As a result, much is known about the
genes carried, the location of restriction sites, etc.
Despite this, the P-group plasmids are not widely
POGC08 9/11/2001 11:06 AM Page 146
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CHAPTER 8
AccI
PvuII
AccI
PvuII
PvuII
PvuII
PvuII
PvuII
tetR
PvuII
lacZ’
SmaI
XmaI
lacZ’
NdeI
tetA
cat
pJB3Tc20
7069 bp
NdeI
pJB3Cm6
6227 bp
trfA
oriT
PvuII
trfA
oriT
AccI
SfiI
Pneo
bla
oriV
PvuII
AccI
SalI
AccI
SfiI
bla
Pneo
oriV
PvuII
AccI
SalI
PstI
PvuII
SmaI
XmaI
HindIII
PvuII
lacZ’
kan
PstI
AccI
SalI
NdeI
trfA
pJB3Km1
6052 bp
oriT
PvuII
PvuII
BgIII
AccI
NdeI
lacZ’ parDE
NdeI
trfA
AccI
pJB321
5594 bp
oriT
bla
SfiI
Pneo
PvuII
bla
SfiI
oriV
Pneo oriV
PvuII
used as vectors because their large size makes
manipulations difficult.
A number of groups have developed mini-IncP
plasmids as vectors and good examples are those
of Blatny et al. (1997). Their vectors are only 4.8–
7.1 kb in size but can still be maintained in a wide
range of Gram-negative bacteria. All the vectors
share a common polylinker and lacZ′ region, thereby
simplifying cloning procedures and identification of
inserts by blue/white screening (see p. 35) and most
carry two antibiotic-resistance determinants. All the
vectors retain the oriT (origin of transfer) locus,
enabling them to be conjugally transferred in those
cases where the recipient cannot be successfully
transformed or electroporated. Two other features
of these vectors deserve mention. First, the parDE
region from the parent plasmid has been included in
some of the vectors, since this greatly enhances their
segregative stability in certain hosts. Secondly, the
trfA locus on the vectors contains unique sites for
the restriction enzymes NdeI and SfiI. Removal of
the NdeI–SfiI fragment results in an increased copy
number. Expression vectors have also been devel-
AccI
Fig. 8.4 Map and construction of
general-purpose broad-host-range
cloning vectors derived from plasmid
RP4. The restriction sites in the
polylinker downstream of the lacZ
promoter are marked (t), and the sites
are, in the counterclockwise direction,
HindIII, SphI, PstI, SalI/HincII/AccI,
XbaI, BamHI, XmaI/SmaI, KpnI, SacI and
EcoRI. Sites in the polylinker that are
not unique are indicated elsewhere
on each vector. Note that the sites for
NdeI and SfiI are unique for all of the
vectors except pJB321. Pneo, promoter
from the neomycin resistance gene;
bla, kan, tet and cat, genes encoding
ampicillin, kanamycin, tetracycline
and chloramphenicol resistance,
respectively. (Figure modified from
Blatny et al. 1997.)
oped by the inclusion of controllable promoters.
Representative examples of these vectors are shown
in Fig. 8.4.
An alternative way of using P-group plasmids as
cloning vectors has been described by Kok et al.
(1994). Their method combines the advantages
of high-copy-number pBR322 vectors with the convenience of conjugative plasmids. This is achieved
by converting the pBR322 vector into a transposable element. Most pBR322 derivatives contain the
β-lactamase gene and one of two 38 bp inverted
repeats of transposon Tn2. By adding a second
inverted repeat, a transposable element is created
(Fig. 8.5). All that is missing is transposase activity
and this is provided by another plasmid, which is a
pSC101 derivative carrying the tnpA gene. To use
this system, the desired DNA sequence is cloned into
the transposition vector. The recombinant molecules
are transformed into an E. coli strain carrying the
P-group plasmid (e.g. R751) and the pSC101 tnpA
derivative and selection is made for the desired characteristics. Once a suitable transformant has been
selected, it is conjugated with other Gram-negative
POGC08 9/11/2001 11:06 AM Page 147
Cloning in bacteria other than E. coli
SstII
HII
Bss R V
Eco I
a
Hp
lacl
lacZ’
aI
Hp oRI
Ec
Bgl II
0
25
lacPO
20
pSPORTn3
10
R
Km
15
Sm
R
Su R
Ap
EcoRI
SstII
pH
Bs
I
pH
Bs
BglII EcoRI
R
Cm
Sal I
SstII
Bgl II
SmaI
I
ScaI
N
aeI
5
pSa
(29.6 kb)
ri
F1 o
BspMI
Sse8387I
PstI
KpnI
AgeI
RsrII
BspEII
EcoRI
SmaI
SalI
SacI
SpeI
NotI
XbaI
BamHI
HindIII
SnaBI
SplI
SphI
AatII
147
Fig. 8.5 A transposable vector derived from pBR322. The
solid arrowheads indicate the inverted repeats required for
transposition. Insertion of a DNA fragment in the multiplecloning site results in inactivation of the lacZα gene and
regulated gene expression from the wild-type lac promoter.
bacteria and selection made for the ampicillinresistance marker carried on the transposon.
Vectors derived from the IncW plasmid Sa
Although a group-W plasmid, such as plasmid pSa
(Fig. 8.6) can infect a wide range of Gram-negative
bacteria, it has been developed mainly as a vector
for use with the oncogenic bacterium A. tumefaciens
(see p. 224). Two regions of the plasmid have been
identified as involved in conjugal transfer of the plasmid and one of them has the unexpected property
of suppressing oncogenicity by A. tumefaciens (Tait
et al. 1982). Information encoding the replication of
the plasmid in E. coli and A. tumefaciens is contained
within a 4 kb DNA fragment. Leemans et al. (1982b)
have described four small (5.6–7.2 MDa), multiply
marked derivatives of pSa. The derivatives contain
single-target sites for a number of the common
restriction endonucleases and at least one marker in
each is subject to insertional inactivation. Although
these Sa derivatives are non-conjugative, they can
be mobilized by other conjugative plasmids. Tait
et al. (1983) have also constructed a set of broadhost-range vectors from pSa. The properties of their
Fig. 8.6 The structure of plasmid Sa. The grey area encodes
the functions essential for plasmid replication. The dark red
areas represent the regions containing functions essential for
self-transmission, the one between the Sst and Sal sites being
responsible for suppression of tumour induction by
Agrobacterium tumefaciens.
derivatives are similar to those of Leemans et al.
(1982b), but one of them also contains the bacteriophage λ cos sequence and hence functions as a
cosmid. Specialist vectors for use in Agrobacterium
and which are derived from a natural Agrobacterium
plasmid have been described by Gallie et al. (1988).
Vectors derived from pBBR1
Plasmid BBR1 is a broad-host-range plasmid originally isolated from Bordatella bronchiseptica that is
compatible with IncP, IncQ and IncW plasmids and
replicates in a wide range of bacteria. Kovach et al.
(1995) have developed a series of vectors from
pBBR1 that are relatively small (< 5.3 kb), possess
multiple cloning sites, allow direct selection of
recombinants in E. coli by blue/white screening and
are mobilizable by IncP plasmids. Newman and
Fuqua (1999) have developed an expression vector
from one of these pBBR1 derivatives by incorporating the araBAD/araC cassette from E. coli (see p. 76)
and shown that the promoter is controllable in
Agrobacterium. Sukchawalit et al. (1999), using a
similar vector, have shown that the promoter is also
controllable in Xanthomonas.
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CHAPTER 8
Cloning in Gram-positive bacteria
In Gram-positive bacteria, the base composition of
the different genomes ranges from < 30% GC to
> 70% GC. Given this disparity in GC content, the
preferred codons and regulatory signals used by
organisms at one end of the % GC spectrum will not
be recognized by organisms at the other end. This
in turn means that there are no universal cloning
vehicles for use with all Gram-positive bacteria.
Rather, one set of systems has been developed for
high-GC organisms (e.g. streptomycetes) and another
for low-GC organisms. This latter group comprises
bacteria from the unrelated genera Bacillus, Clostridium and Staphylococcus and the lactic acid bacteria
Streptococcus, Lactococcus and Lactobacillus.
Vectors for cloning in Bacillus subtilis
and other low-GC organisms
The development of B. subtilis vectors began with
the observation (Ehrlich 1977) that plasmids from
S. aureus (Table 8.1) can be transformed into B.
subtilis, where they replicate and express antibiotic
resistance normally.
As can be seen from Table 8.1, none of the natural
S. aureus plasmids carries more than one selectable
marker and so improved vectors have been constructed by gene manipulation, e.g. pHV11 is pC194
carrying the TcR gene of pT127 (Ehrlich 1978). In
general, these plasmids are stable in B. subtilis, but
segregative stability is greatly reduced following
insertion of exogenous DNA (Bron & Luxen 1985).
Reasoning that stable host–vector systems in B.
subtilis are more likely if endogenous plasmids are
used, Bron and colleagues have developed the cryptic Bacillus plasmid pTA1060 as a vector (Haima
et al. 1987, Bron et al. 1989).
Because of the difficulties experienced in direct
cloning in B. subtilis, hybrid plasmids were constructed which can replicate in both E. coli and
B. subtilis. Originally most of these were constructed
as fusions between pBR322 and pC194 or pUB110.
With such plasmids, E. coli can be used as an efficient
intermediate host for cloning. Plasmid preparations
extracted from E. coli clones are subsequently used
to transform competent B. subtilis cells. Such preparations contain sufficient amounts of multimeric
plasmid molecules to be efficient in B. subtiliscompetent cell transformation (see p. 140).
Table 8.2 lists some of the commonly used shuttle
plasmids. Note that some of them carry some of the
features described earlier for E. coli plasmids, e.g. the
E. coli lacZα-complementation fragment, multiple
cloning sites (MCS) (see p. 53) and the phage f1
origin for subsequent production of single-stranded
DNA in a suitable E. coli host (see p. 70).
The influence of mode of replication:
β1
vectors derived from pAMβ
Early in the development of B. subtilis cloning vectors,
it was noted that only short DNA fragments could be
efficiently cloned (Michel et al. 1980) and that longer
DNA segments often undergo rearrangements (Ehrlich
et al. 1986). This structural instability is independent of the host recombination systems, for it still
occurs in Rec− strains (Peijnenburg et al. 1987).
Table 8.1 Properties of some S. aureus plasmids used as vectors in B. subtilis.
Plasmid
Phenotype conferred
on host cell
Size
Copy no.
Other comments
pC194
Chloramphenicol
resistance
2906 bp
15
Generates large amount of high-molecular-weight DNA when carrying
heterologous inserts
pE194
Erythromycin
resistance
3728 bp
10
cop-6 derivative has copy number of 100.
Plasmid is naturally temperature-sensitive for replication
pUB110
Kanamycin
resistance
4548 bp
50
Virtually the complete sequence is involved in replication maintenance,
site-specific plasmid recombination or conjugal transfer
POGC08 9/11/2001 11:06 AM Page 149
Cloning in bacteria other than E. coli
149
Table 8.2 B. subtilis–E. coli shuttle plasmids.
Replicon
Markers
Size
(kbp)
E. coli
pHV14
4.6
pBR322
pC194
Ap, Cm
Cm
pBR322/pC194 fusion. Sites: PstI, BamHI, SalI, NcoI
(Ehrlich 1978)
pHV15
4.6
pBR322
pC194
Ap, Cm
Cm
pHV14, reversed orientation of pC194 relative to pBR322
pHV33
4.6
pBR322
pC194
Ap, Tc, Cm
Cm
Revertant of pHV14 (Primrose & Ehrlich 1981)
pEB10
8.9
pBR322
pUB110
Ap, Km
Km
pBR322/pUB110 fusion (Bron et al. 1988)
pLB5
5.8
pBR322
pUB110
Ap, Cm, Km
Cm, Km
Deletion of pBR322/pUB110 fusion, CmR gene of pCl94
Segregationally unstable (Bron & Luxen 1985). Sites:
BamHI, EcoRI, BqlIII (in KmR gene), NcoI (in CmR gene)
pHP3
4.8
pBR322
pTA1060
Em, Cm
Em, Cm
Segregationally stable pTA1060 replicon (Peeters et al.
1988). Copy number c. 5. Sites: NcoI (CmR gene), BclI
and HpaI (both EmR gene)
pHP3Ff
5.3
pBR322
pTA1060
Em, Cm
Em, Cm
Like pHP3; phage f1 replication origin and packaging
signal
pGPA14
5.8
pBR322
pTA1060
Em
Em
Stable pTA1060 replicon. Copy number c. 5.
a-Amylase-based selection vector for protein export
functions (Smith et al. 1987). MCS of M13mp11 in lacZa
pGPB14
5.7
pBR322
pTA1060
Em
Em
As pGPA14, probe gene TEM-b-lactamase
pHP13
4.9
pBR322
pTA1060
Em, Cm
Em, Cm
Stable pTA1060 replicon. Copy number c. 5. Efficient
(shotgun) cloning vector (Haima et al. 1987). MCS of
M13mp9 in lacZa
LacZa not expressed in B. subtilis. Additional sites: Bc lI
and HpaI (both EmR gene)
pHV1431
10.9
pBR322
pAMb1
Ap, Tc, Cm
Cm
Efficient cloning vector based on segregationally stable
pAMb1 (Jannière et al. 1990). Copy number c. 200. Sites:
BgmHI, SalI, PstI, NcoI. Structurally unstable in E. coli
pHV1432
8.8
pBR322
pAMb1
Ap, Tc, Cm
Cm
pHV1431 lacking stability fragment orfH. Structurally
stable in E. coli
pHV1436
8.2
pBR322
pTB19
Ap, Tc, Cm
Cm
Low-copy-number cloning vector (Jannière et al. 1990)
Structurally stable
Plasmid
B. subtilis
E. coli
B. subtilis
A major contributing factor to structural instability of recombinant DNA in B. subtilis appears to be
the mode of replication of the plasmid vector (Gruss
& Ehrlich 1989, Jannière et al. 1990). All the B. subtilis vectors described above replicate by a rollingcircle mechanism (see Box 8.2). Nearly every step in
the process digresses or could digress from its usual
function, thus effecting rearrangements. Also, singlestranded DNA is known to be a reactive intermediate
Comments
in every recombination process, and single-stranded
DNA is generated during rolling-circle replication.
If structural instability is a consequence of rollingcircle replication, then vectors which replicate by
the alternative theta mechanism could be more
stable. Jannière et al. (1990) have studied two potentially useful plasmids, pAMβ1 and pTB19, which
are large (26.5 kb) natural plasmids derived from
Streptococcus (Enterococcus) faecalis and B. subtilis,
POGC08 9/11/2001 11:06 AM Page 150
Box 8.2 The two modes of replication of circular DNA molecules
There are two modes of replication of circular DNA
molecules: via theta-like structures or by a rollingcircle type of mechanism. Visualization by electron
microscopy of the replicating intermediates of many
circular DNA molecules reveals that they retain a ring
structure throughout replication. They always possess
a theta-like shape that comes into existence by the
initiation of a replicating bubble at the origin of
replication (Fig. B8.1). Replication can proceed either
uni- or bidirectionally. As long as each chain remains
intact, even minor untwisting of a section of the
circular double helix results in the creation of positive
supercoils in the other direction. This supercoiling is
relaxed by the action of topoisomerases (see Fig. 4.1),
which create single-stranded breaks (relaxed
molecules) and then reseal them.
An alternative way to replicate circular DNA is the
rolling-circle mechanism (Fig. B8.2). DNA synthesis
starts with a cut in one strand at the origin of
replication. The 5’ end of the cut strand is displaced
from the duplex, permitting the addition of
deoxyribonucleotides at the free 3’ end. As
replication proceeds, the 5’ end of the cut strand is
rolled out as a free tail of increasing length. When a
full-length tail is produced, the replicating machinery
cuts it off and ligates the two ends together. The
double-stranded progeny can reinitiate replication,
whereas the single-stranded progeny must first be
converted to a double-stranded form. Gruss and
Ehrlich (1989) have suggested how deletants and
defective molecules can be produced at each step in
the rolling-circle process.
ori
5‘
5‘
+
5‘
Fig. B8.1 Theta replication of a circular DNA molecule.
The original DNA is shown in black and the newlysynthesized DNA in red. l represents the origin of
replication and the arrow shows the direction of
replication.
Fig. B8.2 Rolling-circle replication of a circular DNA
molecule. The original DNA is shown in black and the
newly synthesized DNA in red. The solid and open circles
represent the positions of the replication origins of the
outer (+) and inner (−) circles, respectively.
POGC08 9/11/2001 11:06 AM Page 151
Cloning in bacteria other than E. coli
respectively. Replication of these plasmids does not
lead to accumulation of detectable amounts of singlestranded DNA, whereas the rolling-circle mode
of replication does. Also, the replication regions of
these two large plasmids share no sequence homology with the corresponding highly conserved regions
of the rolling-circle-type plasmids. It is worth noting
that the classical E. coli vectors, which are derived
from plasmid ColE1, all replicate via theta-like
structures.
Renault et al. (1996) have developed a series of
cloning vectors from pAMβ1. All the vectors carry a
gene essential for replication, repE, and its regulator,
copF. The latter gene can be inactivated by inserting
a linker into its unique KpnI site. Since copF downregulates the expression of repE, its inactivation
leads to an increase in the plasmid copy number
per cell. The original low-copy-number state can be
restored by removal of the linker by cleavage and
religation. This new replicon has been used to build
vectors for making transcriptional and translational
fusions and for expression of native proteins. Poyart
and Trieu-Cuot (1997) have constructed a shuttle
vector based on pAMβ1 for the construction of transcriptional fusions; it can be conjugally transferred
between E. coli and a wide range of Gram-positive
bacteria.
Transcription and translation
The composition of the core RNA polymerase in
B. subtilis and other low-GC hosts resembles that of
E. coli. The number of sigma factors is different in each
of the various genera but the principal sigma factor
is sigma A. Analysis of many sigma A-dependent
Bacillus promoters shows that they contain the
canonical −35 and −10 sequences found in E. coli
promoters. In B. subtilis, at least, many promoters contain an essential TGTG motif (−16 region) upstream
of the −10 region. Mutations of this region significantly reduce promoter strength (Helmann 1995,
Voskuil & Chambliss 1998). The promoters also have
conserved polyA and polyT tracts upstream of the
−35 region. Although the −16 region is found in some
E. coli promoters, such promoters often lack the −35
region, whereas this never occurs in B. subtilis.
The translation apparatus of B. subtilis differs
significantly from that of E. coli (for review, see
151
Vellanoweth 1993). This is demonstrated by the
observation that E. coli ribosomes can support protein synthesis when directed by mRNA from a range
of Gram-positive and Gram-negative organisms,
whereas ribosomes from B. subtilis recognize only
homologous mRNA (Stallcup et al. 1974). The
explanation for the selectivity of B. subtilis ribosomes
is that they lack a counterpart of the largest E. coli
ribosomal protein, S1 (Higo et al. 1982, Roberts &
Rabinowitz 1989). Other Gram-positive bacteria,
such as Staphylococcus, Streptococcus, Clostridium and
Lactobacillus, also lack an S1-equivalent protein and
they too exhibit mRNA selectivity. The role of S1 is
believed to be to bind RNA non-specifically and bring
it to the decoding site of the 30S subunit, where proper
positioning of the Shine–Dalgarno (S-D) sequence
and initiation codon signals can take place. This
is reflected in a more extensive complementarity
between the S-D sequences and the 3′ end of the
16S ribosomal RNA (rRNA) than found in bacteria
which have ribosomal protein S1.
The additional sequence requirements for efficient
transcription and translation in B. subtilis and other
low-GC organisms probably explain why many E. coli
genes are not expressed in these hosts.
Controlled expression in B. subtilis and
other low-GC hosts
The first controlled expression system to be used in
B. subtilis was the Spac system (Yansura & Henner
1984). This consists of the E. coli lacI gene and the
promoter of phage SPO-1 coupled to the lac operator.
More recently, the E. coli T7 system (see p. 74) has
been successfully implemented in B. subtilis (Conrad
et al. 1996). This was achieved by inserting the T7
RNA polymerase gene (rpoT7) into the chromosome
under the control of a xylose-inducible promoter
and cloning the gene of interest, coupled to a T7 promoter, on a B. subtilis vector. Of course, expression
of the heterologous gene can be made simpler by
putting it directly under the control of the xyloseinducible promoter (Kim et al. 1996). A similar
xylose-inducible system has been developed in
staphylococci (Sizemore et al. 1991, Peschel et al.
1996) and Lactobacillus (Lokman et al. 1997). Many
different controllable promoters are available in
Lactococcus lactis (for reviews see Kuipers et al. 1995,
POGC08 9/11/2001 11:06 AM Page 152
152
CHAPTER 8
Table 8.3 Some inducible systems in L. lactis.
Promoter
Inducer
lacA or lacR
dnaJ
sodA
PA170
trpE
f31 and ori
nisA or nisF
Lactose
High temperature
Aeration
Low pH, low temperature
Absence of tryptophan
f31 infection
Nisin
De Vos et al. 1997) and some representative examples are shown in Table 8.3.
In the φ31 system of L. lactis, the gene of interest is
placed under the control of a phage middle promoter
inserted in a low-copy-number vector carrying the
phage ori region. Following infection of the host cell
with φ31, the plasmid copy number rapidly increases
and this is followed by expression from the phage
promoter. Following induction in this manner, the
level of expression of the cloned gene can increase
over 1000-fold (O’Sullivan et al. 1996). Similar
levels of expression can be achieved by using the
nisA and nisF systems but with the added advantage
that the exact level of expression depends on the
amount of nisin added to the medium (Kuipers et al.
1995, De Ruyter et al. 1996).
Secretion vectors for low-GC bacteria
The export mechanism in Bacillus and other low-GC
bacteria resembles that of E. coli (for review, see
Tjalsma et al. 2000). However, there are differences
in the signal peptides compared with those found
in E. coli and eukaryotes. The NH2 termini are
more positively charged. The signal peptides are also
larger and the extra length is distributed among all
three regions of the signal peptide. Hols et al. (1992)
developed two probe vectors for the identification
of Gram-positive secretion signals. These vectors
made use of a silent reporter gene encoding the
mature α-amylase from Bacillus licheniformis. The disadvantage of this system is that detection of secreted
amylase involves flooding starch-containing media
with iodine and this kills the bacteria in the colonies.
Consequently, replica plates must be made before
iodine addition.
Poquet et al. (1998) have developed an alternative probe system which uses the S. aureus-secreted
nuclease as a reporter. This nuclease is a small (168
amino acid), stable, monomeric enzyme that is devoid
of cysteine residues and the enzymatic test is nontoxic to bacterial colonies. The probe vectors have the
nuclease gene, lacking its signal sequence, located
downstream from an MCS. Cloning DNA in the vectors results in the synthesis of fusion proteins and
those containing signal sequences are detected by
nuclease activity in the growth medium. Le Loir et al.
(1998) have noted that inclusion of a nine-residue
synthetic propeptide immediately downstream of
the signal-peptide cleavage site significantly enhances
secretion.
Vectors for systematic gene inactivation
With the advent of mass sequencing of genomes
(see p. 2), many genes have been discovered whose
function is unknown. One way of determining
function is to inactivate the gene and then monitor
the effect of this on cell fitness under different growth
conditions. To study the functions of uncharacterized open reading frames in B. subtilis, Vagner et al.
(1998) constructed a series of vectors to perform
directed insertional mutagenesis in the chromosome. These vectors, which have been given the
designation pMUTIN, have the following properties:
• an inability to replicate in B. subtilis, which allows
insertional mutagenesis;
• a reporter lacZ gene to facilitate the measurement
of expression of the target gene;
• the inducible Pspac promoter to allow controlled
expression of genes downstream of and found in the
same operon as the target gene.
A typical pMUTIN vector is shown in Fig. 8.7 and
their mode of use is as follows. An internal fragment
of the target gene is amplified by polymerase chain
reaction (PCR) and cloned in a pMUTIN vector and
the resulting plasmid is used to transform B. subtilis.
Upon integration, the target gene is interrupted and
a transcriptional fusion is generated between its
promoter and the reporter lacZ gene (Fig. 8.8). If the
targeted gene is part of an operon, then any genes
downstream of it are placed under the control of the
POGC08 9/11/2001 11:06 AM Page 153
Cloning in bacteria other than E. coli
Ter
Pspac
MCS
Op
Hind III
EcoRI
NotI
SacII
BamHI
EmR
pMUTIN
lacZ
ApR
ori
lacI
Fig. 8.7 A typical pMUTIN vector. The EmR, lacI and lacZ
genes are expressed in B. subtilis and the ApR gene is expressed
in E. coli. ‘Ter’ indicates the presence of a terminator to
prevent run-through transcription from the EmR gene.
‘Op’ represents the LacI operator.
Pspac promoter. It should be noted that the procedure shown in Fig. 8.8 simultaneously generates
two types of mutants: an absolute (null) mutation in
orf2 through gene inactivation, and a conditional
mutation in orf3, which can be relieved by induction
with isopropyl-β-D-thiogalactoside (IPTG).
153
Cloning in streptomycetes
Cloning in Streptomyces has attracted a lot of interest
because of the large number of antibiotics that are
made by members of this genus. Although Streptomyces coelicolor is the model species for genetic
studies (Hopwood 1999), many other species are the
subject of intensive study and methods developed
for one species may not work particularly well in
another.
Streptomycete DNA has a G+C content of 70–75%
and this affects the frequency of restriction sites.
As might be expected, AT-rich recognition sites are
rare and this can be useful if large-sized fragments
of DNA are wanted. For the construction of gene
libraries, the most commonly used enzymes are
ones with a high GC content in their recognition
sequence, e.g. BamH1 (G′GATCC), BglII (A′GATCT),
and BclI (T′GATCA).
In Streptomyces, promoters may be several hundred base pairs upstream of the start of the gene and
so can be lost during gene cloning. Also, many
Streptomyces promoters are complex and may
include tandem sites for recognition by different
sigma factors. Streptomycetes are good at expressing genes (promoters, ribosome binding sites, etc.)
from low-G+C organisms, but Streptomyces genes
are usually difficult to express in E. coli because most
promoters do not function, and translation may be
inefficient unless the initial amino acid codons are
changed to lower-G+C alternatives.
ori
Ap
lacl
R
EmR
lacZ
Pspac
Fig. 8.8 Integration of pMUTIN into
a target gene. Genes of the orf1–orf3
operon are indicated as white boxes.
Red box corresponds to the internal
segment of the target gene. The vector is
integrated in orf2 by a single crossingover event. (Figure reproduced for
Microbiology courtesy of Dr S.D. Ehrlich
and the Society for General
Microbiology.)
orf1
lacZ
orf2
lacI
ori
orf1
orf3
ApR
∆orf2
EmR
Pspac
orf3
POGC08 9/11/2001 11:06 AM Page 154
154
CHAPTER 8
There are several ways in which DNA can be
introduced into streptomycetes, including transformation, transfection and conjugation. Transformation is achieved by using protoplasts, rather than
competent cells, and high frequencies of plasmid DNA
uptake can be achieved in the presence of polyethylene glycol (Bibb et al. 1978). Plasmid monomers that are covalently closed will yield 106–107
transformants/µg of DNA, even with plasmids up to
60 kb in size. Open circular and linearized molecules
with sticky ends transform with 10–100-fold lower
efficiency (Bibb et al. 1980). The number of transformants obtained with non-replicating plasmids
that integrate by homologous recombination into
the recipient chromosome is greatly stimulated by
simple denaturation of the donor DNA (Oh & Chater
1997). This stimulation reflects an increased frequency of recombination rather than an increased
frequency of DNA uptake. Electroporation has been
used to transform streptomycetes, since it bypasses
the need to develop protoplast regeneration procedures (Pigac & Schrempf 1995, Tyurin et al.
1995). For electroporation, limited non-protoplasting
lysozyme treatment is used to weaken the cell wall
and improve DNA uptake. Intergeneric conjugation
of mobilizable plasmids from E. coli into streptomycetes (see p. 142) is increasingly being used,
because the required constructs can be made easily
in E. coli and the conjugation protocols are simple.
For intergeneric conjugation to occur, the vectors
have to carry the oriT locus from RP4 and the E. coli
strain needs to supply the transfer functions in trans
(Mazodier et al. 1989).
Transformants are generally identified by the
selection of appropriate phenotypes. However, antibiotic resistance has much less utility than in other
organisms, because many streptomycetes produce
antibiotics and hence have innate resistance to
them. One particularly useful phenomenon is that
clones harbouring conjugative plasmids can be
detected by the visualization of pocks. The property
of pock formation, also known as lethal zygosis, is
exhibited if a strain containing a conjugative plasmid
is replica-plated on to a lawn of the corresponding
plasmid-free strain. Under these conditions, clones
containing plasmids are surrounded by a narrow
zone in which the growth of the plasmid-free strain
is retarded (Chater & Hopwood 1983).
Table 8.4 Streptomyces plasmids that have been used
in the development of vectors.
Plasmid
pIJ101
pJV1
pSG5
SCP2*
SLP1
pSAM2
Size
Mode of
replication
8.8 kb
11.1 kb
12.2 kb
31 kb
17.2 kb
10.9 kb
Rolling circle
Rolling circle
Rolling circle
Theta
Rolling circle
Rolling circle
Copy
number
Host
range
300
20–50
1–4
Integrating
Integrating
Broad
Broad
Limited
Broad
Vectors for streptomycetes
With the exception of RSF1010 (see p. 145), no plasmid from any other organism has been found to
replicate in Streptomyces. For this reason, all the
cloning vectors used in streptomycetes are derived
from plasmids and phages that occur naturally in
them. The different replicons that have been subjugated as vectors are listed in Table 8.4. Nearly all
Streptomyces plasmids carry transfer functions that
permit conjugative plasmid transfer and provide
different levels of chromosome-mobilizing activity.
These transfer functions are very simple, consisting
of a single transfer gene and a repressor of gene
function.
Plasmid SCP2* is a deriviative (Lydiate et al. 1985)
of the sex plasmid SCP2. Both plasmids have a size
of 31.4 kb and are physically indistinguishable,
although SCP2* exhibits a much more pronounced
lethal zygosis reaction. SCP2* is important because
it is the progenitor of many very low-copy-number,
stable vectors. High-copy-number derivatives have
also been isolated with the exact copy number (10 or
1000) being dependent on the sequences from the
replication region that are present. SLP1 and pSAM2
are examples of Streptomyces plasmids that normally
reside integrated into a specific highly conserved
chromosomal transfer RNA (tRNA) sequence (Kieser
& Hopwood 1991). Many different specialist vectors
have been derived from these plasmids, including
cosmids, expression vectors, vectors with promoterless reporter genes, positive-selection vectors,
temperature-sensitive vectors, etc., and full details
can be found in Kieser et al. (2000).
POGC08 9/11/2001 11:06 AM Page 155
Cloning in bacteria other than E. coli
The temperate phage φC31 is the streptomycete
equivalent of phage λ and has been subjugated as a
vector. φC31-derived vectors have upper and lower
size limits for clonable fragments with an average
insert size of 8 kb. In contrast, there are no such size
constraints on plasmid cloning, although recombinant plasmids of a size greater than 35 kb are rare with
the usual vectors. However, phage vectors do have
one important advantage: plaques can be obtained
overnight, whereas plasmid transformants can take
up to 1 week to sporulate. Plasmid-integrating vectors
can be generated by incorporating the integration
functions of φC31.
As noted earlier, a major reason for cloning in
streptomycetes is to analyse the genetics and regulation of antibiotic synthesis. Although all the genes
for a few complete biosynthetic pathways have been
cloned (Malpartida & Hopwood 1985, Kao et al.
1994, Schwecke et al. 1995), some gene clusters may
be too large to be cloned in the standard vectors. For
155
this reason, Sosio et al. (2000) generated bacterial
artificial chromosomes (BACs) that can accommodate
up to 100 kb of streptomycete DNA. These vectors
can be shuttled between E. coli, where they replicate autonomously, and Streptomyces, where they
integrate site-specifically into the chromosome.
Homoeologous recombination
Homoeologous recombination is the recombination between DNA sequences that are only partially
homologous. In most bacteria, homoeologous recombination fails to occur because of mismatch
repair, but it does occur in streptomycetes, although
the frequency of recombination is about 105-fold
lower than for homologous recombination. The
significance of homoeologous recombination is that
it permits the formation of hybrid genes, gene clusters or even species and can lead to the formation of
new antibiotics (Baltz 1998).
POGC09 9/11/2001 11:05 AM Page 156
CHAPTER 9
Cloning in Saccharomyces
cerevisiae and other fungi
Introduction
Introducing DNA into fungi
The analysis of eukaryotic DNA sequences has been
facilitated by the ease with which DNA from eukaryotes can be cloned in prokaryotes, using the vectors
described in previous chapters. Such cloned sequences
can be obtained easily in large amounts and can be
altered in vivo by bacterial genetic techniques and in
vitro by specific enzyme modifications. To determine
the effects of these experimentally induced changes
on the function and expression of eukaryotic genes,
the rearranged sequences must be taken out of the
bacteria in which they were cloned and reintroduced
into a eukaryotic organism. Despite the overall unity
of biochemistry, there are many functions common
to eukaryotic cells which are absent from prokaryotes, e.g. localization of ATP-generating systems to
mitochondria, association of DNA with histones,
mitosis and meiosis, and obligate differentiation of
cells. The genetic control of such functions must be
assessed in a eukaryotic environment.
Ideally these eukaryotic genes should be reintroduced into the organism from which they were
obtained. In this chapter we shall discuss the potential for cloning these genes in Saccharomyces cerevisiae and other fungi and in later chapters we shall
consider methods for cloning in animal and plant
cells. It should be borne in mind that yeast cells are
much easier to grow and manipulate than plant and
animal cells. Fortunately, the cellular biochemistry
and regulation of yeast are very like those of higher
eukaryotes. For example, signal transduction and
transcription regulation by mammalian steroid
receptors can be mimicked in strains of S. cerevisiae
expressing receptor sequences (Metzger et al. 1988,
Schena & Yamamoto 1988). There are many yeast
homologues of human genes, e.g. those involved in
cell division. Thus yeast can be a very good surrogate host for studying the structure and function of
eukaryotic gene products.
Like Escherichia coli, fungi are not naturally transformable and artificial means have to be used for
introducing foreign DNA. One method involves the
use of spheroplasts (i.e. wall-less cells) and was first
developed for S. cerevisiae (Hinnen et al. 1978). In
this method, the cell wall is removed enzymically and
the resulting spheroplasts are fused with ethylene
glycol in the presence of DNA and CaCl2. The
spheroplasts are then allowed to generate new cell
walls in a stabilizing medium containing 3% agar.
This latter step makes subsequent retrieval of cells
inconvenient. Electroporation provides a simpler and
more convenient alternative to the use of spheroplasts. Cells transformed by electroporation can be
selected on the surface of solid media, thus facilitating subsequent manipulation. Both the spheroplast
technique and electroporation have been applied to
a wide range of yeasts and filamentous fungi.
DNA can also be introduced into yeasts and
filamentous fungi by conjugation. Heinemann and
Sprague (1989) and Sikorski et al. (1990) found that
enterobacterial plasmids, such as R751 (IncPβ) and
F (IncF), could facilitate plasmid transfer from E. coli
to S. cerevisiae and Schizosaccharomyces pombe. The
bacterial plant pathogen Agrobacterium tumefaciens
contains a large plasmid, the Ti plasmid, and part
of this plasmid (the transfered DNA (T-DNA)) can be
conjugally transferred to protoplasts of S. cerevisiae
(Bundock et al. 1995) and a range of filamentous
fungi (De Groot et al. 1998). T-DNA can also be
transferred to hyphae and conidia.
The fate of DNA introduced into fungi
In the original experiments on transformation of S.
cerevisiae, Hinnen et al. (1978) transformed a leucine
auxotroph with the plasmid pYeLeu 10. This plasmid
is a hybrid composed of the enterobacterial plasmid
POGC09 9/11/2001 11:05 AM Page 157
Cloning in S. cerevisiae and other fungi
ColE1 and a segment of yeast DNA containing the
LEU2+ gene and is unable to replicate in yeast.
Analysis of the transformants showed that in some
of them there had been reciprocal recombination
between the incoming LEU2+ and the recipient
Leu2− alleles. In the majority of the transformants,
ColE1 DNA was also present and genetic analysis
showed that in some of them the LEU2+ allele was
closely linked to the original Leu2− allele, whereas
in the remaining ones the LEU2+ allele was on a
different chromosome.
The results described above can be confirmed by
restriction-endonuclease analysis, since pYeLeu 10
contains no cleavage sites for HindIII. When DNA
from the Leu2− parent was digested with endonuclease HindIII and electrophoresed in agarose, multiple
DNA fragments were observed but only one of these
hybridized with DNA from pYeLeu 10. With the
transformants in which the Leu2− and LEU2+ alleles
were linked, only a single fragment of DNA hybridized
to pYeLeu 10, but this had an increased size, consistent with the insertion of a complete pYeLeu 10
molecule into the original fragment. These data are
consistent with there being a tandem duplication of
the Leu2 region of the chromosome (Fig. 9.1). With
the remaining transformants, two DNA fragments
that hybridized to pYeLeu 10 could be found on
electrophoresis. One fragment corresponded to the
fragment seen with DNA from the recipient cells,
the other to the plasmid genome which had been
157
inserted in another chromosome (see Fig. 10.1). These
results represented the first unambiguous demonstration that foreign DNA, in this case cloned ColE1
DNA, can integrate into the genome of a eukaryote.
A plasmid such as pYeLeu 10 which can do this is
known as a yeast integrating plasmid (YIp).
During transformation, the integration of exogenous DNA can occur by recombination with a homologous or an unrelated sequence. In most cases,
non-homologous integration is more common than
homologous recombination (Fincham 1989), but
this is not so in S. cerevisiae (Schiestl & Petes 1991).
In the experiments of Hinnen et al. (1978) described
above, sequences of the yeast retrotransposon Ty2
were probably responsible for the integration of
the plasmid in novel locations of the genome, i.e.
the ‘illegitimate’ recombinants were the result of
homologous crossovers within a repeated element
(Kudla & Nicolas 1992). Based on a similar principle, a novel vector has been constructed by Kudla
and Nicolas (1992) which allows integration of
a cloned DNA sequence at different sites in the
genome. This feature is provided by the inclusion
in the vector of a repeated yeast sigma sequence
present in approximately 20–30 copies per genome
and spread over most or all of the 16 chromosomes.
When T-DNA from the Ti plasmid of Agrobacterium
is transferred to yeast, it too will insert in different
parts of the genome by illegitimate recombination
(Bundock & Hooykaas 1996).
Chromosome structure of
transformants and recipient
Electrophoretic separation
of HindIII generated
fragments which hybridize
with pYeleu 10
HindIII HindIII
Recipient
Leu 2–
Leu 2+
Transformants
Leu 2– pYeleu10
Leu 2–
pYeleu10
Fig. 9.1 Analysis of yeast transformants.
(See text for details.)
Direction of migration
POGC09 9/11/2001 11:05 AM Page 158
CHAPTER 9
Xba I
Ase I
Ahd I
a
SnaB I
2µ
cir
mp
Hpa I
A
If the heterologous DNA introduced into fungi is to
be maintained in an extrachromosomal state then
plasmid vectors are required which are capable of
replicating in the fungal host. Four types of plasmid
vector have been developed: yeast episomal plasmids
(YEps), yeast replicating plasmids (YRps), yeast
centromere plasmids (YCps) and yeast artificial
chromosomes (YACs). All of them have features in
common. First, they all contain unique target sites
for a number of restriction endonucleases. Secondly,
they can all replicate in E. coli, often at high copy
number. This is important, because for many experiments it is necessary to amplify the vector DNA in
E. coli before transformation of the ultimate yeast
recipient. Finally, they all employ markers that
can be selected readily in yeast and which will often
complement the corresponding mutations in E. coli
as well. The four most widely used markers are His3,
Leu2, Trp1 and Ura3. Mutations in the cognate chromosomal markers are recessive, and non-reverting
mutants are available. Two yeast selectable markers, Ura3 and Lys2, have the advantage of offering
both positive and negative selection. Positive selection is for complementation of auxotrophy. Negative
selection is for ability to grow on medium containing
a compound that inhibits the growth of cells expressing the wild-type function. In the case of Ura3,
it is 5-fluoro-orotic acid (Boeke et al. 1984) and
for Lys2 it is α-aminoadipate (Chatoo et al. 1979).
These inhibitors permit the ready selection of those
rare cells which have undergone a recombination or
loss event to remove the plasmid DNA sequences.
The Lys2 gene is not utilized frequently because it is
Mfe I
DN
Plasmid vectors for use in fungi
Bcl I
Aat II
cle
Schiestl and Petes (1991) developed a method for
forcing illegitimate recombination by transforming
yeast with BamH1-generated fragments in the presence of the BamH1 enzyme. Not only did this increase
the frequency of transformants but the transformants
which were obtained had the exogenous DNA integrated into genomic BamH1 sites. This technique,
which is sometimes referred to as restriction-enzymemediated integration (REMI), has been extended to
other fungi, such as Cochliobolus (Lu et al. 1994),
Ustilago (Bolker et al. 1995) and Aspergillus (Sanchez
et al. 1998).
ORI
158
Blp I
YEp 24
Tth111I
Cla I -BspD I
Pvu II
BsaB I
BspE I
a3
Sbf I
ur
Tc
Eag I
PshA I
Sal I
EcoN I
Sph I
Nco I
Api I/Bsp1201 I
Sma I/Xma I
Nhe I
BamH I
SgrA I
Fig. 9.2 Schematic representation of a typical yeast episomal
plasmid (YEp 24). The plasmid can replicate both in E. coli
(due to the presence of the pBR322 origin of replication) and
in S. cerevisiae (due to the presence of the yeast 2 µm origin of
replication). The ampicillin and tetracycline determinants
are derived from pBR322 and the URA3 gene from yeast.
large and contains sites within the coding sequence
for many of the commonly used restriction sites.
Yeast episomal plasmids
YEps were first constructed by Beggs (1978) by
recombining an E. coli cloning vector with the naturally occurring yeast 2 µm plasmid. This plasmid is
6.3 kb in size, has a copy number of 50–100 per
haploid cell and has no known function. A representative YEp is shown in Fig. 9.2.
Yeast replicating plasmids
YRps were initially constructed by Struhl et al. (1979).
They isolated chromosomal fragments of DNA which
carry sequences that enable E. coli vectors to replicate in yeast cells. Such sequences are known as
ars (autonomously replicating sequence). An ars is
quite different from a centromere: the former acts
as an origin of replication (Palzkill & Newlon 1988,
Huang & Kowalski 1993), whereas the latter is
involved in chromosome segregation.
POGC09 9/11/2001 11:05 AM Page 159
Cloning in S. cerevisiae and other fungi
Although plasmids containing an ars transform
yeast very efficiently, the resulting transformants
are exceedingly unstable. For unknown reasons, YRps
tend to remain associated with the mother cell and
are not efficiently distributed to the daughter cell.
(Note: S. cerevisiae does not undergo binary fission
but buds off daughter cells instead.) Occasional
stable transformants are found and these appear to
be cases in which the entire YRp has integrated into
a homologous region on a chromosome in a manner
identical to that of YIps (Stinchcomb et al. 1979,
Nasmyth & Reed 1980).
Yeast centromere plasmids
Using a YRp vector, Clarke and Carbon (1980)
isolated a number of hybrid plasmids containing
DNA segments from around the centromere-linked
leu2, cdc10 and pgk loci on chromosome III of yeast.
As expected for plasmids carrying an ars, most of the
recombinants were unstable in yeast. However, one
of them was maintained stably through mitosis
and meiosis. The stability segment was confined to a
1.6 kb region lying between the leu2 and cdc10 loci
and its presence on plasmids carrying either of two
ars tested resulted in those plasmids behaving like
minichromosomes (Clarke & Carbon 1980, Hsiao &
Carbon 1981). Genetic markers on the minichromosomes acted as linked markers segregating in the
first meiotic division as centromere-linked genes and
were unlinked to genes on other chromosomes.
Structurally, plasmid-borne centromere sequences
have the same distinctive chromatin structure that
occurs in the centromere region of yeast chromosomes (Bloom & Carbon 1982). Functionally YCps
exhibit three characteristics of chromosomes in
yeast cells. First, they are mitotically stable in the
absence of selective pressure. Secondly, they segregate during meiosis in a Mendelian manner. Finally,
they are found at low copy number in the host cell.
Yeast artificial chromosomes
All three autonomous plasmid vectors described above
are maintained in yeast as circular DNA molecules
– even the YCp vectors, which possess yeast centromeres. Thus, none of these vectors resembles the
normal yeast chromosomes which have a linear
159
structure. The ends of all yeast chromosomes, like
those of all other linear eukaryotic chromosomes,
have unique structures that are called telomeres.
Telomere structure has evolved as a device to preserve the integrity of the ends of DNA molecules,
which often cannot be finished by the conventional
mechanisms of DNA replication (for detailed discussion see Watson 1972). Szostak and Blackburn
(1982) developed the first vector which could be
maintained as a linear molecule, thereby mimicking
a chromosome, by cloning yeast telomeres into a
YRp. Such vectors are known as yeast artificial chromosomes (YACs).
One advantage of YACs is that, unlike the other
plasmid vectors, their stability increases as the
size of the insert increases. Thus, there is no practical limitation to the size of a YAC and they are
essential tools in any genome-sequencing project.
The method for cloning large DNA sequences in
YACs developed by Burke et al. (1987) is shown in
Fig. 9.3.
Retrovirus-like vectors
The genome of S. cerevisiae contains 30– 40 copies
of a 5.9 kb mobile genetic element called Ty (for
review see Fulton et al. 1987). This transposable
element shares many structural and functional
features with retroviruses (see p. 193) and the
copia element of Drosophila. Ty consists of a central
region containing two long open reading frames
(ORFs) flanked by two identical terminal 334 bp
repeats called delta (Fig. 9.4). Each delta element
contains a promoter as well as sequences recognized
by the transposing enzyme. New copies of the transposon arise by a replicative process, in which the Ty
transcript is converted to a progeny DNA molecule
by a Ty-encoded reverse transcriptase. The complementary DNA can transpose to many sites in the
host DNA.
The Ty element has been modified in vitro by
replacing its delta promoter sequence with promoters derived from the phosphoglycerate kinase
or galactose-utilization genes (Garfinkel et al. 1985,
Mellor et al. 1985). When such constraints are introduced into yeast on high-copy-number vectors, the
Ty element is overexpressed. This results in the
formation of large numbers of virus-like particles
POGC09 9/11/2001 11:05 AM Page 160
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CHAPTER 9
Target DNA
EcoRI
cloning site
CEN4
ARS1
URA 3
TRP1
pYAC vector
AMP
ori
TEL
TEL
BamHI
BamHI
BamHI and
EcoRI-digest
Partial
EcoRI-digest
TRP ARS CEN
URA
Ligate
TEL
TRP ARS CEN
URA
LTR
LTR
Primary transcript
ORF 1
ORF 2
Fig. 9.4 Structure of a typical Ty element. ORF 1 and ORF 2
represent the two open reading frames. The delta sequences
are indicated by LTR (long terminal repeats).
(VLPs), which accumulate in the cytoplasm (Fig. 9.5).
The particles, which have a diameter of 60–80 nm,
have reverse-transcriptase activity. The major structural components of VLPs are proteins produced
by proteolysis of the primary translation product of
TEL
Fig. 9.3 Construction of a yeast
artificial chromosome containing large
pieces of cloned DNA. Key regions of the
pYAC vector are as follows: TEL, yeast
telomeres; ARS 1, autonomously
replicating sequence; CEN 4,
centromere from chromosome 4; URA3
and TRP1, yeast marker genes; Amp,
ampicillin-resistance determinant of
pBR322; ori, origin of replication of
pBR322.
ORF 1. Adams et al. (1987) have shown that fusion
proteins can be produced in cells by inserting part of
a gene from human immunodeficiency virus (HIV)
into ORF 1. Such fusion proteins formed hybrid
HIV:Ty-VLPs.
The Ty element can also be subjugated as a vector
for transposing genes to new sites in the genome.
The gene to be transposed is placed between the 3′
end of ORF 2 and the 3′ delta sequence (Fig. 9.6).
Providing the inserted gene lacks transcriptiontermination signals, transcription of the 3′ delta
sequence will occur, which is a prerequisite for
transposition. Such constructs act as amplification
cassettes, for, once introduced into yeast, transposition of the new gene occurs to multiple sites in the
genome (Boeke et al. 1988, Jacobs et al. 1988).
POGC09 9/11/2001 11:05 AM Page 161
Cloning in S. cerevisiae and other fungi
161
Fig. 9.5 Ty virus-like particles (magnification 80 000) carrying the entire HIV1 TAT coding region. (Photograph courtesy of
Dr S. Kingsman.)
δ
pGAL
2µ ORI
Ty transcript
P
Transcript
of gene
insert
δ
URA 3
Fig. 9.6 Structure of the multicopy plasmid used for inserting
a modified Ty element, carrying a cloned gene, into the yeast
chromosome. pGAL and P are yeast promoters, δ represents
the long terminal repeats (delta sequences) and the red region
represents the cloned gene. (See text for details.)
Choice of vector for cloning
There are three reasons for cloning genes in yeast.
The first of these relates to the potential use of yeast
as a cloning host for the overproduction of proteins of
commercial value. Yeast offers a number of advantages, such as the ability to glycosylate proteins during secretion and the absence of pyrogenic toxins.
Commercial production demands overproduction and
the factors affecting expression of genes in yeast are
discussed in a later section (see p. 165). Yeast is also
used in the production of food and beverages. The
ability to clone in yeast without the introduction of
bacterial sequences by using vectors like those of Chinery and Hinchliffe (1989) is particularly beneficial.
A second reason for cloning genes in yeast is the
ability to clone large pieces of DNA. Although there
is no theoretical limit to the size of DNA which can
be cloned in a bacterial plasmid, large recombinant
plasmids exhibit structural and segregative instability. In the case of bacteriophage-λ vectors, the size of
the insert is governed by packaging constraints. Many
DNA sequences of interest are much larger than this.
For example, the gene for blood Factor VIII covers
about 190 kbp, or about 0.1% of the human X chromosome, and the Duchenne muscular dystrophy gene
spans more than a megabase. Long sequences of
cloned DNA have greatly facilitated efforts to sequence
the human genome. YACs offer a convenient way
to clone large DNA fragments but are being replaced
with BACs and PACs (see p. 67) because these have
greater stability. Nevertheless, the availability of
YACs with large inserts was an essential prerequisite
for the early genome-sequencing projects and they
were used in the sequencing of the entire S. cerevisiae
chromosome III (Oliver et al. 1992). The method for
POGC09 9/11/2001 11:05 AM Page 162
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CHAPTER 9
Table 9.1 Properties of the different yeast vectors.
Vector
Transformation
frequency
Copy
no./cell
Loss in
non-selective
medium
Disadvantages
Advantages
Much less than
1% per
generation
1 Low transformation
frequency
2 Can only be recovered
from yeast by cutting
chromosomal DNA with
restriction endonuclease
which does not cleave
original vector
containing cloned gene
1 Of all vectors, this kind give most
stable maintenance of cloned genes
2 An integrated YIp plasmid behaves
as an ordinary genetic marker, e.g.
a diploid heterozygous for an
integrated plasmid segregates the
plasmid in a Mendelian fashion
3 Most useful for surrogate genetics
of yeast, e.g. can be used to
introduce deletions, inversions and
transpositions (see Botstein &
Davis 1982)
1% per
generation
Novel recombinants
generated in vivo by
recombination with
endogenous 2 mm
plasmid
1 Readily recovered from yeast
2 High copy number
3 High transformation frequency
4 Very useful for complementation
studies
YIp
102 transformants
per mg DNA
YEp
103–105
transformants
per mg DNA
YRp
104 transformants
per mg DNA
1–20
Much greater
than 1% per
generation but
can get
chromosomal
integration
Instability of
transformants
1 Readily recovered from yeast
2 High copy number. Note that the
copy number is usually less than that
of YEp vectors but this may be useful
if cloning gene whose product is
deleterious to the cell if produced in
excess
3 High transformation frequency
4 Very useful for complementation
studies
5 Can integrate into the chromosome
YCp
104 transformants
per mg DNA
1–2
Less than 1%
per generation
Low copy number makes
recovery from yeast
more difficult than that
with YEp or YRp vectors
1 Low copy number is useful if
product of cloned gene is deleterious
to cell
2 High transformation frequency
3 Very useful for complementation
studies
4 At meiosis generally shows
Mendelian segregation
1–2
Depends on
length: the
longer the YAC
the more
stable it is
Difficult to map by
standard techniques
1 High-capacity cloning system
permitting DNA molecules greater
than 40 kb to be cloned
2 Can amplify large DNA molecules
in a simple genetic background
Stable, since
integrated into
chromosome
Needs to be introduced
into cell in another
vector
Can get amplification following
chromosomal integration
YAC
Ty
Depends on
vector used to
introduce Ty
into cell
1
25–200
~20
POGC09 9/11/2001 11:05 AM Page 163
Cloning in S. cerevisiae and other fungi
cloning large DNA sequences developed by Burke et
al. (1987) is shown in Fig. 9.3).
For many biologists the primary purpose of cloning is to understand what particular genes do in vivo.
Thus most of the applications of yeast vectors have
been in the surrogate genetics of yeast. One advantage of cloned genes is that they can be analysed easily,
particularly with the advent of DNA-sequencing
methods. Thus nucleotide-sequencing analysis can
reveal many of the elements that control expression
of a gene, as well as identifying the sequence of
the gene product. In the case of the yeast actin gene
(Gallwitz & Sures 1980, Ng & Abelson 1980) and
some yeast transfer RNA (tRNA) genes (Peebles et al.
1979, Olson 1981), this kind of analysis revealed the
presence within these genes of non-coding sequences
which are copied into primary transcripts. These
introns are subsequently eliminated by a process
known as splicing. Nucleotide-sequence analysis can
also reveal the direction of transcription of a gene,
although this can be determined in vivo by other
methods. For example, if the yeast gene is expressed
in E. coli using bacterial transcription signals, the
direction of reading can be deduced by observing the
orientation of a cloned fragment required to permit
expression. Finally, if a single transcribed yeast gene
is present on a vector, the chimera can be used as a
probe for quantitative solution hybridization analysis
of transcription of the gene.
The availability of different kinds of vectors with
different properties (see Table 9.1) enables yeast
geneticists to perform manipulations in yeast like
those long available to E. coli geneticists with their
sex factors and transducing phages. Thus cloned
genes can be used in conventional genetic analysis
by means of recombination using YIp vectors or
linearized YRp vectors (Orr-Weaver et al. 1981).
Complementation can be carried out using YEp,
YRp, YCp or YAC vectors, but there are a number of
factors which make YCps the vectors of choice (Rose
et al. 1987). For example, YEps and YRps exist at
high copy number in yeast and this can prevent the
isolation of genes whose products are toxic when
overexpressed, e.g. the genes for actin and tubulin.
In other cases, the overexpression of genes other
than the gene of interest can suppress the mutation
used for selection (Kuo & Campbell 1983). All the
yeast vectors can be used to create partial diploids or
163
partial polyploids and the extra gene sequences can
be integrated or extrachromosomal. Deletions, point
mutations and frame-shift mutations can be introduced in vitro into cloned genes and the altered genes
returned to yeast and used to replace the wild-type
allele. Excellent reviews of these techniques have
been presented by Botstein and Davis (1982), Hicks
et al. (1982), Struhl (1983) and Stearns et al. (1990).
Plasmid construction by homologous
recombination in yeast
During the process of analysing a particular cloned
gene it is often necessary to change the plasmid’s
selective marker. Alternatively, it may be desired
to move the cloned gene to a different plasmid, e.g.
from a YCp to a YEp. Again, genetic analysis may
require many different alleles of a cloned gene to be
introduced to a particular plasmid for subsequent
functional studies. All these objectives can be achieved by standard in vitro techniques, but Ma et al.
(1987) have shown that methods based on recombination in vivo are much quicker. The underlying
principle is that linearized plasmids are efficiently
repaired during yeast transformation by recombination with a homologous DNA restriction fragment.
Suppose we wish to move the HIS3 gene from
pBR328, which cannot replicate in yeast, to YEp420
(see Fig. 9.7). Plasmid pRB328 is cut with PvuI
and PvuII and the HIS3 fragment selected. The HIS3
fragment is mixed with YEp420 which has been
linearized with EcoRI and the mixture transformed
into yeast. Two crossover events occurring between
homologous regions flanking the EcoRI site of YEp420
will result in the generation of a recombinant YEp
containing both the HIS3 and URA3 genes. The
HIS3 gene can be selected directly. If this were not
possible, selection could be made for the URA3 gene,
for a very high proportion of the clones will also
carry the HIS3 gene.
Many other variations of the above method have
been described by Ma et al. (1987), to whom the
interested reader is referred for details.
Expression of cloned genes
When gene manipulation in fungi first became
possible, there were many unsuccessful attempts to
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CHAPTER 9
Initiator
PvuI
UAS
PvuII
URS
Coding
sequence
TATA
AmpR
mRNA
pBR328
40–120 bp
Eco
H IS 3
20–40 bp
Sal
Bam
100–1400 bp
PvuI + PvuII
Fig. 9.8 Structure of typical yeast promoters.
(See text for details.)
Eco
H IS 3
Bam
Sal
PvuI
Bam
Sal
PvuI
PvuII
UR
A3
Eco-cut
YEp420
2µ
Recombination
Eco
PvuI
AmpR
S
HI
3
Bam
Sal
2µ
3
Recombinant
UR
A
Fig. 9.7 Plasmid construction by homologous recombination
in yeast. pRB328 is digested with PvuI and PvuII and the
HIS3-containing fragment is transformed into yeast along
with the EcoRI-cut YEp420. Homologous recombination
occurs between pBR322 sequences, shown as thin lines, to
generate a new plasmid carrying both HIS3 and URA3.
express heterologous genes from bacteria or higher
eukaryotes. This suggested that fungal promoters
have a unique structure, a feature first shown for
S. cerevisiae (Guarente 1987). Four structural elements
can be recognized in the average yeast promoter
(Fig. 9.8). First, several consensus sequences are
found at the transcription-initiation site. Two of these
sequences, TC(G/A)A and PuPuPyPuPu, account
for more than half of the known yeast initiation
sites (Hahn et al. 1985, Rudolph & Hinnen 1987).
These sequences are not found at transcriptioninitiation sites in higher eukaryotes, which implies
a mechanistic difference in their transcription
machinery compared with yeast.
The second motif in the yeast promoter is the
TATA box (Dobson et al. 1982). This is an AT-rich
region with the canonical sequence TATAT/AAT/A,
located 60–120 nucleotides before the initiation
site. Functionally, it can be considered equivalent to
the Pribnow box of E. coli promoters (see p. 51).
The third and fourth structural elements are
upstream activating sequences (UASs) and upstream
repressing sequences (URSs). These are found in
genes whose transcription is regulated. Binding of
positive-control proteins to UASs increases the rate
of transcription and deletion of the UASs abolishes
transcription. An important structural feature of
UASs is the presence of one or more regions of
dyad symmetry (Rudolph & Hinnen 1987). Binding
of negative-control proteins to URSs reduces the
transcription rate of those genes that need to be
negatively regulated.
The level of transcription can be affected by
sequences located within the gene itself and which
are referred to as downstream activating sequences
(DASs). Chen et al. (1984) noted that, when using
the phosphoglycerate kinase (PGK ) promoter, several heterologous proteins accumulate to 1–2% of
total cell protein, whereas phosphoglycerate kinase
itself accumulates to over 50%. These disappointing
amounts of heterologous protein reflect the levels of
POGC09 9/11/2001 11:05 AM Page 165
Cloning in S. cerevisiae and other fungi
mRNA which were due to a lower level of initiation
rather than a reduced mRNA half-life (Mellor et al.
1987). Addition of downstream PGK sequences
restored the rate of mRNA transcription, indicating
the presence of a DAS. Evidence for these DASs has
been found in a number of other genes.
Overexpression of proteins in fungi
The first overexpression systems developed were
for S. cerevisiae and used promoters from genes
165
encoding abundant glycolytic enzymes, e.g. alcohol
dehydrogenase (ADH1), PGK or glyceraldehyde-3phosphate dehydrogenase (GAP). These are strong
promoters and mRNA transcribed from them can
accumulate up to 5% of total. They were at first
thought to be constitutive but later were shown to
be induced by glucose (Tuite et al. 1982). Now there
is a large variety of native and engineered promoters
available (Table 9.2), differing in strength, regulation
and induction ratio. These have been reviewed in
detail by Romanos et al. (1992).
Table 9.2 Common fungal promoters used for manipulation of gene expression.
Species
General
S. cerevisiae
Candida albicans
Methanol utilizers
Candida boidnii
Hansenula polymorpha
Pichia methanolica
Pichia pastoris
Lactose utilizer
Kluyveromyces lactis
Starch utilizer
Schwanniomyces
occidentalis
Xylose utilizer
Pichia stipitis
Alkane utilizer
Yarrowia lipolytica
Promoter
Gene
Regulation
PGK
GAL1
PHOS
ADH2
CUP1
MFa1
Phosphoglycerate kinase
Galactokinase
Acid phosphatase
Alcohol dehydrogenase II
Copper metallothionein
Mating factor a1
MET3
ATP sulphur lyase
AOD1
MOX
AUG1
AOX1
GAP
FLD1
PEX8
YPT1
Alcohol oxidase
Alcohol oxidase
Alcohol oxidase
Alcohol oxidase
Glyceraldehyde-3-phosphate
dehydrogenase
Formaldehyde dehydrogenase
Peroxin
Secretory GTPase
Methanol- or methylamine-induced
Methanol-induced
Medium constitutive
LAC4
PGK
ADH4
b-Galactosidase
Phosphoglycerate kinase
Alcohol dehydrogenase
Lactose-induced
Strong constitutive
Ethanol-induced
AMY1
a-Amylase
Maltose- or starch-induced
GAM1
Glucoamylase
Maltose- or starch-induced
XYL1
XPR2
TEF
RPS7
Glucose-induced
Galactose-induced
Phosphate-repressed
Glucose-repressed
Copper-induced
Constitutive but temperature-induced
variant available
Repressed by methionine and cysteine
Methanol-induced
Methanol-induced
Methanol-induced
Methanol-induced
Strong constitutive
Xylose-induced
Extracellular protease
Translation elongation factor
Ribosomal protein S7
Peptone-induced
Strong constitutive
Strong constitutive
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CHAPTER 9
The ideal promoter is one that is tightly regulated
so that the growth phase can be separated from
the induction phase. This minimizes the selection of
non-expressing cells and can permit the expression
of proteins normally toxic to the cell. The ideal
promoter will also have a high induction ratio. One
promoter which has these characteristics and which
is now the most widely used is that from the GAL1
gene. Galactose regulation in yeast is now extremely
well studied and has become a model system for
eukaryotic transcriptional regulation (see Box 9.1).
Following addition of galactose, GAL1 mRNA is
rapidly induced over 1000-fold and can reach 1%
of total mRNA. However, the promoter is strongly
repressed by glucose and so in glucose-grown cultures this induction only occurs following depletion
of glucose. To facilitate galactose induction in the
presence of glucose, mutants have been isolated which
are insensitive to glucose repression (Matsumoto
et al. 1983, Horland et al. 1989). The trans-activator
GAL4 protein is present in only one or two molecules
per cell and so GAL1 transcription is limited. With
multicopy expression vectors, GAL4 limitation is
exacerbated. However, GAL4 expression can be made
autocatalytic by fusing the GAL4 gene to a GAL10
promoter (Schultz et al. 1987), i.e. GAL4 expression
is now regulated (induced) by galactose.
In recent years, methylotrophic yeasts, such as
Pichia pastoris, have proved extremely popular as
hosts for the overexpression of heterologous proteins. There are a number of reasons for this. First,
the alcohol oxidase (AOX1) promoter is one of the
strongest and most regulatable promoters known.
Second, it is possible to stably integrate expression
plasmids at specific sites in the genome in either
single or multiple copies. Third, the strains can be
cultivated to very high density. To date, over 300
foreign proteins have been produced in P. pastoris
(Cereghino & Cregg 1999, 2000). Promoters for use
in other yeasts are shown in Table 9.2.
Specialist vectors
Many different specialist yeast vectors have been
developed which incorporate the useful features
found in the corresponding E. coli vectors (see
p. 70), e.g. an f1 origin to permit sequencing of
inserts production of the cloned gene product as a
purification fusion, etc. Some representative examples are shown in Fig. 9.9. Two aspects of these
vectors warrant further discussion: secretion and
surface display.
In yeast, proteins destined for the cell surface or
for export from the cell are synthesized on and
translocated into the endoplasmic reticulum. From
there they are transported to the Golgi body for
processing and packaging into secretory vesicles.
Fusion of the secretory vesicles with the plasma
membrane then occurs constitutively or in response
to an external signal (reviewed by Rothman & Orci
1992). Of the proteins naturally synthesized and
secreted by yeast, only a few end up in the growth
medium, e.g. the mating pheromone α factor and
the killer toxin. The remainder, such as invertase
and acid phosphatase, cross the plasma membrane
but remain within the periplasmic space or become
associated with the cell wall.
Polypeptides destined for secretion have a
hydrophobic amino-terminal extension, which is
responsible for translocation to the endoplasmic
reticulum (Blobel & Dobberstein 1975). The extension is usually composed of about 20 amino acids
and is cleaved from the mature protein within the
endoplasmic reticulum. Such signal sequences precede the mature yeast invertase and acid phosphatase sequences. Rather longer leader sequences
precede the mature forms of the α mating factor
and the killer toxin (Kurjan & Herskowitz 1982,
Bostian et al. 1984). The initial 20 amino acids or so
are similar to the conventional hydrophobic signal
sequences, but cleavage does not occur in the endoplasmic reticulum. In the case of α factor, which has
an 89 amino acid leader sequence, the first cleavage
occurs after a Lys–Arg sequence at positions 84 and
85 and happens in the Golgi body ( Julius et al. 1983,
1984).
To date, a large number of non-yeast polypeptides
have been secreted from yeast cells containing the
appropriate recombinant plasmid and in almost all
cases the α-factor signal sequence has been used.
There is a perception that S. cerevisiae has a lower
secretory capacity than P. pastoris and other yeasts
(Muller et al. 1998), but the real issue may be the
type of vector used. For example, Parekh et al.
(1996) found that S. cerevisiae strains containing
one stably integrated copy of an expression cassette
POGC09 9/11/2001 05:24 PM Page 167
Box 9.1 Galactose metabolism and its control in Saccharomyces cerevisiae
Galactose is metabolized to glucose-6-phosphate in
yeast by an identical pathway to that operating in
other organisms (Fig. B9.1). The key anzymes and
their corresponding genes are a kinase (GAL1), a
transferase (GAL7), an epimerase (GAL10) and a
mutase (GAL5). Melibiose (galactosyl-glucose) is
metabolized by the same enzymes after cleavage
by an α-galactosidase encoded by the MEL1 gene.
Galactose uptake by yeast cells is via a permease
encoded by the GAL2 gene. The GAL5 gene is
constitutively expressed. All the others are induced
by growth on galactose and repressed during growth
on glucose.
The GAL1, GAL7 and GAL10 genes are clustered
on chromosome II but transcribed separately from
individual promoters. The GAL2 and MEL1 genes are
on other chromosomes. The GAL4 gene encodes a
protein that activates transcription of the catabolic
genes by binding UAS 5′ to each gene. The GAL80
gene encodes a repressor that binds directly to GAL4
gene product, thus preventing it from activating
transcription. The GAL3 gene product catalyses
the conversion of galactose to an inducer, which
combines with the GAL80 gene product, preventing
it from inhibiting the GAL4 protein from binding to
DNA (Fig. B9.2).
The expression of the GAL genes is repressed
during growth on glucose. The regulatory circuit
responsible for this phenomenon, termed catabolite
repression, is superimposed upon the circuit
responsible for induction of GAL gene expression.
Very little is known about its mechanism.
For a review of galactose metabolism in S. cerevisiae,
the reader should consult Johnston (1987).
Medium
MEL1
α-galactosidase
Melibiose
Galactose
Cytoplasm
GAL2
Permease
GAL1
Kinase
Galactose
GAL7
Transferase
GAL5
Mutase
Gal–1–P
Glu–1–P
UDP–Glu
UDP–Gal
Glu–6–P
Epimerase
GAL10
Fig. B9.1 The genes and enzymes associated with the metabolism of galactose by yeast.
GAL3
Galactose
Inducer
GAL80 protein–inducer
complex
GAL4 protein binds to UASs
and activates GAL transcription
GAL80
protein
No
inducer
Fig. B9.2 The regulation of transcription of the yeast galactose genes.
GAL80 protein complexes
GAL4 protein and prevents
activation of GAL transcription
Glycolysis
POGC09 9/11/2001 11:05 AM Page 168
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CHAPTER 9
6xHis
STOP
BstB I
EcoR I
Pml I
Sfi I
BsmB I
Asp718 I
Kpn I
Xho I
Sac II
Not I
Apa I*
Xba I*
SnaB I*
V5 epitope
pPICZ A, B, C
Pme I
T7
Hind III
Asp718 I
Kpn I
Sac I
BamH I
BstX I
EcoR I
BstX I
Not I
Xho I
Xba I*
Polylinkers for C-terminal tagged vectors
P GAL
CYC1
1
c-myc epitope
AOX1
pA
TT
STOP
BamH I
pPICZ A, B, C
and
pPICZα A, B, C
3.3 kb
P EM7
Zeoc
i
n
Blasticid
in
Amplici
llin
4
SH
3
F1
5’ A
OX1
ori
f1
o
ri
C
/AR
in
rig
6
CEN
o
2µ
URA
6xHis
P TE
pU
YES
Vectors
STOP
Pme I
STOP
Cla I*
Pst I*
EcoR I
Pml I
Sfi I
BsmB I
Asp718 I
Kpn I
Xho I
Sac II
Not I
Xba I
Asp718 I
Kpn I
BamH I
BstX I
EcoR I
BstX I
Not I
Xho I
Xba I
Hind III
α-factor
ATG 6xHis Xpress Epitope EK Site V5 epitope 6xHis
6xHis
pPICZα A, B, C
Polylinkers for N-terminal tagged vectors
T7
c-myc epitope
Bgl II
ColE
1
1
CYC
TT
TRP1
Fig. 9.9 Two specialized vectors for use in Saccharomyces (YES vectors) and Pichia (pPICZ). V5, Express, 6XHis and c-myc encode
epitopes which can be readily detected and purified by affinity chromatography. The YES vectors offer a choice of 2 µm origin for
high copy or CEN6/ARSH4 origin for low copy in yeast, a choice of URA3 or TRP1 genes for auxotrophic selection on minimal
medium, blasticidin resistance for dominant selection in any strain and an f1 origin to facilitate DNA sequencing. In the pPICZ
vectors, the zeocin-resistance gene is driven by the EM-7 promoter for selection in E. coli and the TEF1 promoter for selection in
Pichia. More details of these features can be found in Chapter 5. (Figure reproduced courtesy of Invitrogen Corporation.)
secreted more bovine pancreatic trypsin inhibitor
than strains with the same expression cassette on a
2 µm multicopy vector. Optimal expression was
obtained with 10 integrated copies. With those
proteins which tend to accumulate in the endoplasmic reticulum as denatured aggregates, secretion
may be enhanced by simultaneously overexpressing
chaperons (Shusta et al. 1998). Secretion may also
be enhanced by minor amino acid changes. Katakura
et al. (1999) noted a sixfold increase in lactoglobulin
secretion by conversion of a tryptophan residue to
tyrosine.
Yeast surface display
S. cerevisiae can be used to elucidate and dissect the
function of a protein in a manner similar to phagedisplay systems. Either can be used to detect protein–
ligand interactions and to select mutant proteins
with altered binding capacity (Shusta et al. 1999).
However, phage-display systems often cannot display secreted eukaryotic proteins in their native
functional conformation, whereas yeast surface
display can.
Yeast surface display makes use of the cell surface
receptor α-agglutinin (Aga), which is a two-subunit
glycoprotein. The 725-residue Aga1p subunit anchors
the assembly to the cell wall via a covalent linkage.
The 69-residue binding subunit (Aga2p) is linked to
Aga1p by two disulphide bonds. To achieve surface
display, the appropriate gene is inserted at the C
terminus of a vector-borne Aga2p gene under the
control of the GAL1 promoter. The construct is then
transformed into a yeast strain carrying a chromosomal copy of the Aga1p gene, also under the
control of the GAL1 promoter. If the cloned gene has
POGC09 9/11/2001 11:05 AM Page 169
Cloning in S. cerevisiae and other fungi
c-myc
(a)
Cloned
gene
product
169
Aga 2
HA
(b)
MCS
c-myc
Aga2 cassette
R
A1
T
HA
PG
P
TRP1
s
s
s
N/
s
ARS
Aga 2
i
been inserted in the correct translational reading
frame, its gene product will be synthesized as a
fusion with the Aga2p subunit. The fusion product
will associate with the Aga1p subunit within the
secretory pathway and be exported to the cell surface (Boder & Wittrup 1997).
In practice, the gene fusion is somewhat more
complicated. Usually the cloned gene product is
sandwiched between two simple epitopes to permit
quantitation of the number of fusion proteins per
cell and to determine the accessibility of different
domains of the fusion protein (Fig. 9.10).
CE
Aga 1
or
Fig. 9.10 Yeast surface display of
heterologous proteins. (a) Schematic
representation of surface display.
(b) A vector used for facilitating surface
display. MCS, multiple cloning site;
HA, haemagglutinin epitope; c-myc,
c-myc epitope. See text for details.
Yeast cell wall
R
n
Ampicilli
DNA-binding domain hybrid
BAIT
DBD
GAL4 (1–147)
UASG
GAL1–lacZ
Activation domain hybrids encoded by a library
PREY
Y1-n
TAD
GAL4 (768–881)
Detecting protein–protein interactions
Chien et al. (1991) have made use of the properties of
the GAL4 protein to develop a method for detecting
protein–protein interactions. The GAL4 protein has
separate domains for the binding to UAS DNA and
for transcriptional activation. Plasmids were constructed which encode two hybrid proteins. The first
consisted of the DNA-binding domain (residues
1–147) of the GAL4 protein fused to a test protein.
The second consisted of residues 768–881 of the
GAL4 protein, representing the activation domain,
fused to protein sequences encoded by a library of
yeast genomic DNA fragments. Interaction between
the test protein (bait) and a protein encoded by one
of the library plasmids (prey) led to transcriptional
activation of a reporter gene (Fig. 9.11). This method
is known as the two-hybrid system. The two-hybrid
system has become a major tool in the study of
protein–protein and protein–ligand interactions (see
UASG
GAL1–lacZ
Interaction between DNA-binding domain
hybrid and a hybrid from the library
BAIT
DBD
UASG
PREY
Yi
TAD
GAL4 (768–881)
GAL4 (1–147)
GAL1–lacZ
Fig. 9.11 Strategy to detect interacting proteins using
the two-hybrid system. UASG is the upstream activating
sequence for the yeast GAL genes, which binds the GAL4
protein, DBD is a DNA-binding domain and TAD is a
transcription-activation domain. Interaction is detected by
expression of a GAL1– lacZ gene fusion. (Reproduced courtesy
of Dr S. Fields and the National Academy of Sciences.)
POGC09 9/11/2001 11:05 AM Page 170
Box 9.2 Variations of the two-hybrid system
of the bait with a previously identified partner
protein may permit expression of the reporter
gene ( Tomashek et al. 1996). The ternary partner
in such three-component interactions need
not be a protein but can be a small molecule,
such as a hormone (Lee et al. 1995) or a drug
(Chiu et al. 1994).
Post-translational modifications
A limitation of using the two-hybrid system is that
certain protein–protein interactions are not possible
in S. cerevisiae because of the absence of the
appropriate post-translational modifying enzymes.
This is a particular problem with baits relevant to
signal transduction in higher eukaryotes, because
of the prevalence of site-specific phosphorylation
by tyrosine kinase as a means of regulating protein–
protein interactions. Osborne et al. (1996) were able
to solve this problem by using a strain of yeast
expressing the mammalian tyrosine kinase Lck.
Bait and hook
The two-hybrid system can be modified to study
the interaction between proteins and RNA
(SenGupta et al. 1999) or proteins and drugs
(Licitra & Liu 1996). To do this a ‘hook’ is used to
display an RNA molecule or a drug to the incoming
prey, thereby forming a non-protein bridge to the
reporter gene (Fig. B9.3).
Protein–peptide interactions
The utility of the two-hybrid system can be extended
to interactions between proteins and very small
(< 16 residues) peptides (Yang et al. 1995, Colas
et al. 1996). Small peptides can be used not only
as the prey, as in the studies just cited, but also as
the bait (Geyer et al. 1999, Norman et al. 1999).
In another variation, Stagljar et al. (1996) have
described a rapid method to identify small
interaction-specific sequences within larger proteins.
In their method, complementary DNA (cDNA)
encoding a known interacting protein is sheared
by sonication and shotgun-cloned into a prey vector.
The resulting targeted library is cotransformed
with the bait of interest and direct selection made
for fragments capable of conferring the desired
interaction.
Reverse two-hybrid system
The original two-hybrid system relies on
transcriptional activation of reporter genes to
detect protein–protein interactions (Fig. 9.11).
However, the two-hybrid system can be ‘turned on its
head’ to monitor loss of protein–protein interactions.
For example, Vidal et al. (1996a,b), have described a
screen for mutations that disrupted the ability of the
transcription factor E2F1 to associate with its partner
DP1. In the absence of interaction between the two
partner proteins, transcription of the URA3 reporter
gene was abolished. This in turn allowed the host
yeasts to grow in the presence of 5-fluoro-orotic acid,
a compound that is toxic to cells capable of
synthesizing uracil. This type of reverse selection
can be used to screen for peptides or other small
molecules that disrupt protein–protein interactions
and that could have utility as drugs (Huang &
Schreiber 1997).
Three-component systems
In some cases, a bait and prey may not directly
interact, or else form a transient contact, because a
third component is missing. However, co-expression
Hook
Bait RNA
Prey
DBD
UAS
TAD
Transcription
Reporter
Fig. B9.3 The bait-and-hook
variation of the two-hybrid system.
POGC09 9/11/2001 11:05 AM Page 171
Cloning in S. cerevisiae and other fungi
DBD1
Fig. 9.12 The dual-bait two-hybrid
system. The first DNA-binding domain
fusion protein (DBD1–Bait1) drives
the expression of reporter 1 through
UAS1. A second separate DNA-binding
domain fused to a distinct bait
(DBD2–Bait2) directs expression of
reporter 2 through UAS2. In this
example, the prey can interact with
Bait 1 but not Bait 2.
Box 9.2 and the reviews by Colas & Brent 1998, and
Uetz & Hughes 2000).
A major operational problem with the two-hybrid
system has been the high frequency of false positives,
i.e. cells in which the reporter gene is active even
though the bait and prey do not interact in nature
and/or in yeast. There are two general solutions to
this problem. In the first of these ( James et al. 1996),
a host strain is used that contains three reporter
genes under the control of different GAL4-inducible
promoters. An alternative approach is to use the
two-bait system (Fig. 9.12). In this system, the yeast
expresses a single prey and two independent baits,
each targeted to distinct reporter genes (Xu et al.
1997). In weeding out false positives, one looks for
preys that interact with only one bait. The rationale
here is that the more related the two baits are, the
more likely it is that a prey interacting with only one
of them is a natural interaction.
Identifying genes encoding particular
cellular activities
As a result of a major genome-sequencing project,
the sequences of genes encoding every biochemical
activity of S. cerevisiae are known. The next task is
to connect specific biochemical activities with particular genes. Martzen et al. (1999) have developed
a rapid and sensitive method for doing this which
is applicable to almost any activity. The startingpoint was a yeast vector containing the glutathioneS-transferase (GST) gene under the control of the
CUP1 promoter. A large number of different DNA
fragments bearing yeast ORFs were cloned in this
vector to generate GST–ORF fusions. In all, 6144
Bait1
171
Prey
TAD
UAS1
Reporter 1
Prey
DBD2
UAS2
Transcription
TAD
Bait2
Reporter 2
constructs were made and these were transformed
back into S. cerevisiae.
To correlate genes with activity, the different transformants were dispensed into 64 96-well microtitre
plates (64 × 96 = 6144). Initially, all the clones from
each plate were pooled and the 64 pools analysed for
a specific biochemical activity. Analysis is facilitated
since the GST–ORF fusions are readily purified from
other cellular activities by affinity chromatography
using immobilized glutathione. Once a plate was
identified as having the desired activity, pools of
clones were made from each of the eight rows and
12 columns of wells and these 20 pools reassayed. If
only one clone expresses the desired activity, then one
row and one column should be identified (Fig. 9.13)
and the point of intersection identifies the wanted
clone. Sequencing of the ORF then allows identification of the gene corresponding to the activity.
Determining functions associated
with particular genes
Determining the function of a particular gene product
usually involves determining the expression profile of
the gene, the subcellular location of the protein, and
the phenotype of a null strain lacking the protein.
Conditional alleles of the gene are often created as
an additional tool. These procedures usually require
multiple independent manipulations. Ross-Macdonald
et al. (1997) have developed a multifunctional,
transposon-based system that simultaneously generates constructs for all of the above analyses and is
suitable for mutagenesis of any given S. cerevisiae
gene. The transposons used by them are shown in
Fig. 9.14. Each carries a reporter gene (β-galactosidase
POGC09 9/11/2001 11:05 AM Page 172
172
CHAPTER 9
64 pools
each with
96 GST–ORF
fusions
1 positive pool
1 positive row and
1 positive column
Fig. 9.13 Method for selecting
glutathione-S-transferase openreading-frame fusions. (Figure
reproduced from Trends in Genetics
courtesy of Elsevier Press.)
1 positive strain
TR loxR
HA
lacZ
URA3
tet
loxP
TR
res 3xHa
+cre
mTn-4xHA/lacZ
Xa
mTn-3xHA/lacZ
GFP
+cre
mTn-3xHA/GFP
or green fluorescent protein) lacking a promoter.
In-frame fusions between a yeast coding region and
the transposon can be detected by β-galactosidase
activity or fluorescence. The transposon insertion
creates a truncation of the gene, thereby generating
a null phenotype. The method is also adaptable
to genome-wide analyses, using a high-throughput
screening methodology (Ross-Macdonald et al. 1999).
262-bp
HAT tag
274-bp
HAT tag
Fig. 9.14 Schematic representation of
the transposons, and the derived HAT
tag elements, for use in yeast. Each
transposon carries the tetracyclineresistance determinant, a functional
yeast URA3 gene, and the res element
from transposon Tn3. The function
of the res element is to resolve
transposition cointegrates. (Figure
reproduced courtesy of Dr Michael
Snyder and the National Academy of
Sciences.)
To use this transposon system, yeast genes are
cloned in an E. coli strain that overexpresses the Tn3
transposase. The transposon, carried on a derivative
of the sex factor F, is introduced by mating and transposition ensues. Yeast DNA containing the transposon
is excised from the E. coli vector and transformed back
into yeast, with selection made for URA3 activity. Two
types of transformants are obtained. If homologous
POGC09 9/11/2001 11:05 AM Page 173
Cloning in S. cerevisiae and other fungi
(a) PCR deletion cassette
45 bp
KanR
bc
bc
45 bp
bc
45 bp
(b) Homologous recombination
45 bp
KanR
bc
Yeast ORF
Yeast ORF
(c) Heterozygous yeast strain
KanR
bc
bc
Yeast ORF
(d) Haploid yeast strain
bc
KanR
bc
(e) Homozygous diploid yeast strain
bc
KanR
bc
bc
KanR
bc
Fig. 9.15 ‘Knockout’ strain collection. (a) The bar-coded
deletion cassette is generated in two rounds of PCR. It
contains a kanamycin-resistance gene for selection in yeast
and two molecular bar-codes 20 bp long that are unique to
each deletion. The molecular bar-codes are flanked by
common sequence elements that allow for the PCR
amplification of all the bar-codes in a population of yeast
strains. At the end of the cassette are 45 bp sequences that
are homologous to sequences flanking the gene of interest.
(b) The deletion cassette is transformed into diploid yeast
strains. The 45 bp of yeast sequences allows one copy of the
gene of interest to be replaced with the deletion cassette via
homologous recombination. The resulting heterozygous
strains (c) can then be sporulated and dissected to generate
a collection of haploid mutant strains, if viable (d).
Homozygous mutant strains (e) are made by the selective
matings of haploid segregants. (Figure reproduced courtesy
of Elsevier Science Ltd and the authors.)
173
recombination occurs at the URA3 locus, then no
reporter activity is seen. If integration occurs at the
locus of the gene carrying the transposon, reporter
activity is retained but gene function is lost (Fig. 9.14).
This reporter activity can be used to study the expression of the cloned gene under different growth conditions. If the transformants also carry a phage P1 cre
gene (see p. 68), under the control of the GAL1 promoter, excision of most of the transposon sequences
can be induced by growth of the host strain in the
presence of galactose. In most instances, this results
in restoration of gene activity and the gene product
can be purified by affinity purification, using antibodies to the haemagglutinin determinants.
An alternative approach to the large-scale mutational analysis of the yeast genome is to individually
delete each gene using a polymerase chain reaction
(PCR)-based methodology (Winzeler et al. 1999).
For each gene, a deletion cassette is constructed
(Fig. 9.15) that contains a kanamycin resistance
gene, two ‘molecular bar-codes’ and yeast sequences
homologous to the upstream and downstream flanking sequences. This construct is amplified by PCR and
transformed into a diploid yeast strain, where the deletion cassette replaces one of the two chromosomal
gene sequences. The diploid strains are then allowed
to sporulate and those haploid segregants that are
viable are recovered.
The molecular bar-codes are particularly important
since they are unique to each deletion strain within
a pool of many different strains (Shoemaker et al.
1996). The bar-code can be amplified by PCR, using
common primer sequences that flank the bar-code.
A pool of yeast mutant strains can thus be analysed
in a single experiment by amplifying the bar-codes
and probing a DNA microarray (p. 116) containing
sequences complementary to the bar-codes. Variations
on this technique have been used in other organisms
to identify key functions associated with particular
activities. For example, signature-tagged mutagenesis
has been used to identify genes essential for virulence
(for review, see Lehoux & Levesque 2000).
The application of genome-wide mutagenesis techniques to the understanding of the ways in which
the different components of the yeast cell interact
has been reviewed by Delneri et al. (2001) and Vidan
and Snyder (2001).
POGC10 9/11/2001 11:04 AM Page 174
CH A P T E R 10
Gene transfer to animal cells
Introduction
Overview of gene-transfer strategies
Gene transfer to animal cells has been practised now
for over 40 years. Techniques are available for the
introduction of DNA into many different cell types in
culture, either to study gene function and regulation
or to produce large amounts of recombinant protein.
Animal cells are advantageous for the production of
recombinant animal proteins because they perform
authentic post-translational modifications that are
not carried out by bacterial cells and fungi. Cell cultures
have therefore been used on a commercial scale to
synthesize products such as antibodies, hormones,
growth factors, cytokines and viral coat proteins for
immunization. There has been intense research into
the development of efficient vector systems and
transformation methods for animal cells. Although
this research has focused mainly on the use of mammalian cell lines, other systems have also become
popular, such as the baculovirus expression system,
which is used in insects. More recently, research has
focused on the introduction of DNA into animal cells
in vivo. The most important application of this technology is in vivo gene therapy, i.e. the introduction of
DNA into the cells of live animals in order to treat
disease. Viral gene-delivery vectors are favoured for
therapeutic applications because of their efficiency,
but safety concerns have prompted research into
alternative DNA-mediated transfer procedures.
This chapter focuses on the introduction of DNA
into somatic cells. Unlike the situation in plants,
most animal cells are restricted in terms of their
developmental potential and cannot be used to
generate transgenic animals. Mouse embryonic stem
cells (ES cells) are exceptional in this respect because
they are derived from the early embryo prior to the
formation of the germ line. Gene transfer to ES cells is
therefore discussed in the next chapter, along with
the introduction of DNA into oocytes, eggs and cells
of the early embryo.
Gene transfer to animal cells can be achieved essentially via three routes. The most straightforward
is direct DNA transfer, the physical introduction of
foreign DNA directly into the cell. For example, in
cultured cells this can be done by microinjection,
whereas for cells in vivo direct transfer is often
achieved by bombardment with tiny DNA-coated
metal particles. The second route is termed transfection, and this encompasses a number of techniques,
some chemical and some physical, which can be
used to persuade cells to take up DNA from their surroundings. The third is to package the DNA inside an
animal virus, since viruses have evolved mechanisms to naturally infect cells and introduce their
own nucleic acid. The transfer of foreign DNA into a
cell by this route is termed transduction. Whichever
route is chosen, the result is transformation, i.e. a
change of the recipient cell’s genotype caused by the
acquired foreign DNA, the transgene. Transformation can be transient or stable, depending on how
long the foreign DNA persists in the cell.
DNA-mediated transformation
Transformation techniques
DNA/calcium phosphate coprecipitate method
The ability of mammalian cells to take up exogenously supplied DNA from their culture medium was
first reported by Szybalska and Szybalski (1962).
They used total uncloned genomic DNA to transfect
human cells deficient for the enzyme hypoxanthineguanine phosphoribosyltransferase (HPRT). Rare
HPRT-positive cells, which had presumably taken
up fragments of DNA containing the functional
gene, were identified by selection on HAT medium
(p. 177). At this time, the actual mechanism of DNA
POGC10 9/11/2001 11:04 AM Page 175
Gene transfer to animal cells
uptake was not understood. Much later, it was
appreciated that successful DNA transfer in such
experiments was dependent on the formation of a
fine DNA/calcium phosphate coprecipitate, which
first settles on to the cells and is then internalized.
The precipitate must be formed freshly at the time of
transfection. It is thought that small granules of
calcium phosphate associated with DNA are taken
up by endocytosis and transported to the nucleus,
where some DNA escapes and can be expressed
(Orrantia & Chang, 1990). The technique became
generally accepted after its application, by Graham
and Van der Erb (1973), to the analysis of the infectivity of adenoviral DNA. It is now established as a
general method for the introduction of DNA into a
wide range of cell types in culture. However, since
the precipitate must coat the cells, this method is
suitable only for cells growing in monolayers, not
those growing in suspension or as clumps.
As originally described, calcium phosphate transfection was limited by the variable and rather low
proportion of cells that took up DNA (1–2%). Only a
small number of these would be stably transformed.
Improvements to the method have increased the
transfection frequency to 20% for some cell lines
(Chu & Sharp 1981). A variant of the technique,
using a different buffer system, allows the precipitate
to form slowly over a number of hours, and this
can increase stable transformation efficiency by up
to 100-fold when using high-quality plasmid DNA
(Chen & Okayama 1987, 1988).
Other chemical transfection methods
The calcium phosphate method is applicable to many
cell types, but some cell lines are adversely affected
by the coprecipitate due to its toxicity and are hence
difficult to transfect. Alternative chemical transfection methods have been developed to address this
problem. One such method utilizes diethylaminoethyl dextran (DEAE-dextran), a soluble polycationic
carbohydrate that promotes interactions between
DNA and the endocytotic machinery of the cell. This
technique was initially devised to introduce viral
DNA into cells (McCutchan & Pango 1968) but was
later adapted as a method for plasmid DNA transfer
(Milman & Herzberg 1981). The efficiency of the
original procedure was improved by Lopata et al.
175
(1984) and Sussman and Milman (1984) by adding
after-treatments, such as osmotic shock or exposure to chloroquine, the latter having been shown
to inhibit the acidification of endosomal vesicles
(Luthmann & Magnusson 1983). Although efficient
for the transient transfection of many cell types,
DEAE-dextran cannot be used to generate stably
transformed cell lines. Another polycationic chemical, the detergent Polybrene, has been used for
the transfection of Chinese hamster ovary (CHO)
cells, which are not amenable to calcium phosphate
transfection (Chaney et al. 1986).
Phospholipids as gene-delivery vehicles
An alternative to chemical transfection procedures
is to package DNA inside a phosopholipid vesicle,
which interacts with the target cell membrane and
facilitates DNA uptake. The first example of this
approach was provided by Schaffner (1980), who
used bacterial protoplasts containing plasmids to
transfer DNA into mammalian cells. Briefly, bacterial cells were transformed with a suitable plasmid
vector and then treated with chloramphenicol to
amplify the plasmid copy number. Lysozyme was
used to remove the cell walls, and the resulting protoplasts were gently centrifuged on to a monolayer
of mammalian cells and induced to fuse with them,
using polyethylene glycol. A similar strategy was
employed by Wiberg et al. (1987), who used the
haemoglobin-free ghosts of erythrocytes as delivery
vehicles. The procedures are very efficient in terms of
the number of transformants obtained, but they are
also labour-intensive and so have not been widely
adopted as a general transfection method. However,
an important advantage is that they are gentle,
allowing the transfer of large DNA fragments without shearing. Yeast cells with the cell wall removed
(sphaeroplasts) have therefore been used to introduce yeast artificial chromosome (YAC) DNA into
mouse ES cells by this method, for the production of
YAC transgenic mice (see Chapter 11).
More widespread use has been made of artificial
phospholipid vesicles, which are called liposomes
(Schaefer-Ridder et al. 1982). Initial liposome-based
procedures were hampered by the difficulty encountered in encapsulating the DNA, and provided a
transfection efficiency no better than the calcium
POGC10 9/11/2001 11:04 AM Page 176
176
CHAPTER 10
phosphate method. However, a breakthrough came
with the discovery that cationic/neutral lipid mixtures can spontaneously form stable complexes
with DNA that interact productively with the cell
membrane, resulting in DNA uptake by endocytosis
(Felgner et al. 1987, 1994). This low-toxicity transfection method, commonly known as lipofection, is
one of the simplest to perform and is applicable to
many cell types that are difficult to transfect by other
means, including cells growing in suspension (e.g.
Ruysscharet et al. 1994). This technique is suitable
for transient and stable transformation, and is sufficiently gentle to be used with YACs and other large
DNA fragments. The efficiency is also much higher
than that of chemical transfection methods – up to
90% of cells in a culture dish can be transfected. A
large number of different lipid mixtures is available,
varying in efficiency for different cell lines. A unique
benefit of liposome gene-delivery vehicles is their
ability to transform cells in live animals following
injection into target tissues or even the bloodstream.
Transfection efficiency has been improved and targeting to specific cell types achieved by combining
liposomes with viral proteins that promote cell
fusion, nuclear targeting signals and various molecular conjugates that recognize specific cell surface
molecules. The development of liposomes for gene
therapy has been comprehensively reviewed (Scheule
& Cheng 1996, Tseng & Huang 1998).
Electroporation
Electroporation involves the generation of transient,
nanometre-sized pores in the cell membrane, by
exposing cells to a brief pulse of electricity. DNA
enters the cell through these pores and is transported to the nucleus. This technique was first
applied to animal cells by Wong and Neumann
(1982), who successfully introduced plasmid DNA
into mouse fibroblasts. The electroporation technique has been adapted to many other cell types
(Potter et al. 1984). The most critical parameters are
the intensity and duration of the electric pulse, and
these must be determined empirically for different
cell types. However, once optimal electroporation
parameters have been established, the method is
simple to carry out and highly reproducible. The
technique has high input costs, because a specialized
capacitor discharge machine is required that can
accurately control pulse length and amplitude
(Potter 1988). Additionally, larger numbers of cells
may be required than for other methods because,
in many cases, the most efficient electroporation
occurs when there is up to 50% cell death.
In an alternative method, pores are created using
a finely focused laser beam (Kurata et al. 1986).
Although very efficient (up to 0.5% stable transformation), this technique is applicable to only small numbers of cells and has not gained widespread use.
Direct transfer methods
A final group of methods considered in this section
encompasses those in which the DNA is transferred
directly into the cell nucleus. One such procedure is
microinjection, a technique that is guaranteed to
generate successful hits on target cells but that can
only be applied to a few cells in any one experiment.
This technique has been applied to cultured cells
that are recalcitrant to other transfection methods
(Capecchi 1980), but its principal use is to introduce
DNA and other molecules into large cells, such as
oocytes, eggs and the cells of early embryos, as
discussed in Chapter 11. Particle bombardment is
another direct delivery method, initially developed
for the transformation of plants (Chapter 12). This
involves coating small metal particles with DNA and
then accelerating them into target tissues using a
powerful force, such as a blast of high-pressure gas
or an electric discharge through a water droplet.
In animals, this technique is most often used to
transfect multiple cells in tissue slices rather than
cultured cells (e.g. Arnold et al. 1994, Lo et al. 1994).
It has also been used to transfer DNA into skin cells
in vivo (Haynes et al. 1996).
Transformation with
non-replicating DNA
Transient and stable transformation
DNA-mediated transformation of animal cells occurs
in two stages, the first involving the introduction of
DNA into the cell (the transfection stage) and the
second involving its incorporation into the genome
(the integration stage). Transfection is much more
POGC10 9/11/2001 11:04 AM Page 177
Gene transfer to animal cells
efficient than integration; hence a large proportion
of transfected cells never integrate the foreign DNA
they contain. The DNA is maintained in the nucleus
in an extrachromosomal state and, assuming it does
not contain an origin of replication that functions in
the host cell, it persists for just a short time before it is
diluted and degraded. This is known as transient
transformation (the term transient transfection is also
used), reflecting the fact that the properties of the cell
are changed by the introduced transgene, but only
for a short duration. In a small proportion of transfected cells, the DNA will integrate into the genome,
forming a new genetic locus that will be inherited by
all clonal descendants. This is known as stable transformation, and results in the formation of a ‘cell line’
carrying and expressing the transgene. Since integration is such an inefficient process, the rare stably
transformed cells must be isolated from the large
background of non-transformed and transiently
transformed cells by selection, as discussed below.
While stable transformation is required for longterm analytical experiments and the production of
large amounts of recombinant protein over a prolonged time, transient transformation is sufficient
for many types of short-term experiments, such as
determining the efficiency of promoter sequences
attached to a reporter gene (Box 10.1).
Cotransformation and selection of
stable transformants
Following the general acceptance of the calcium
phosphate transfection method, it was shown that
mouse cells deficient for the enzyme thymidine kinase
(TK) could be stably transformed to a wild-type phenotype by transfecting them with the herpes simplex
virus (HSV) Tk gene (Wigler et al. 1977). As for the
HPRT + transformants discussed earlier in the chapter, cells positive for TK can be selected on HAT
medium. This is because both enzymes are required
for nucleotide biosynthesis via the salvage pathway
(Fig. 10.1). In mammals, nucleotides are produced
via two alternative routes, the de novo and salvage
pathways. In the de novo pathway, nucleotides are
synthesized from basic precursors, such as sugars
and amino acids, while the salvage pathway recycles nucleotides from DNA and RNA. If the de novo
pathway is blocked, nucleotide synthesis becomes
177
dependent on the salvage pathway, and this can be
exploited for the selection of cells carrying functional
Hprt and Tk genes. The drug aminopterin blocks the
de novo synthesis of both inosine monophosphate
(IMP) and thymidine monophosphate (TMP) by
inhibiting key enzymes in the de novo pathway. Cells
exposed to animopterin can thus survive only if they
have functional Hprt and Tk genes and a source of
hypoxanthine and thymidine. Hprt+ and Tk+ transformants can therefore both be selected using HAT
medium, which contains hypoxanthine, aminopterin
and thymidine.
In these early experiments, the transgene of
interest conferred a selectable phenotype on the
cell. However, most genes do not generate a conveniently selectable phenotype, and the isolation of
transformants in such experiments was initially
problematic. A breakthrough was made when it was
discovered that transfection with two physically
unlinked DNAs resulted in cotransformation, i.e. the
integration of both transgenes into the genome
(Wigler et al. 1979). To obtain cotransformants,
cultured cells were transfected with the HSV Tk gene
and a vast excess of well-defined DNA, such as the
plasmid pBR322. Cells selected on HAT medium
were then tested by Southern blot hybridization for
the presence of the non-selected plasmid DNA.
Wigler and colleagues found evidence for the presence of non-selected DNA in nearly 90% of the TK+
cells, indicating that the HSV Tk gene could be used
as a selectable marker. In subsequent experiments,
it was shown that the initially unlinked donor
DNA fragments were incorporated into large concatemeric structures up to 2 Mbp in length prior to
integration (Perucho et al. 1980b).
Other selectable markers for animal cells
The phenomenon of cotransformation allows the
stable introduction of any foreign DNA sequence
into mammalian cells as long as a selectable marker
is introduced at the same time. The HSV Tk gene is
representative of a class of genes known as endogenous markers, because they confer a property that is
already present in wild-type cells. A number of such
markers have been used, all of which act in redundant metabolic pathways (Table 10.1). The major
disadvantage of endogenous markers is that they
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CHAPTER 10
Salvage
pathway
Salvaged bases
and nucleosides
De novo
pathway
Nucleotides
De novo source
APRT
Adenine
AK
AMP
Purines
Adenosine
(Aminopterin, methotrexate)
Inosine
DHFR
Hypoxanthine
Xanthine
HPRT
IMP
IMPDH (Mycophenolic acid)
XGPRT (bacterial)
Guanine
XMP
HPRT
GMP
Pyrimadines
CTP
Thymidine
Deoxyuracil
Glycine +
deoxyribose
TK
TK
UTP
TMP
DNFR
(Aminopterin,
methotrexate)
dUMP
CAD
UDP
Aspartate +
deoxyribose
(PALA)
Fig. 10.1 Simplified representation of the de novo and salvage nucleotide synthesis pathways. Top panel: purine synthesis.
De novo purine nucleotide synthesis (shown on the right) initially involves the formation of inosine monophosphate (IMP)
which is then converted into either adenosine monophosphate (AMP) or, via xanthine monophosphate (XMP), guanosine
monophosphate (GMP). The de novo synthesis of IMP requires the enzyme dihydrofolate reductase (DHFR), whose activity can
be blocked by aminopterin or methotrexate. In the presence of such inhibitors, cell survival depends on nucleotide salvage, as
shown on the left. Cells lacking one of the essential salvage enzymes, such as HPRT or APRT, therefore cannot survive in the
presence of aminopterin or methotrexate unless they are transformed with a functional copy of the corresponding gene. Thus,
the genes encoding salvage-pathway enzymes can be used as selectable markers. Note that the enzyme XGPTR, which converts
xanthine to XMP, is found only in bacterial cells and not in animals. Bottom panel: pyrimidine synthesis. De novo pyrimidine
nucleotide synthesis (shown on the right) initially involves the formation of uridine diphosphate (UDP). This step requires a
multifunctional enzyme, CAD, whose activity can be blocked by N-phosphonacetyl-l-aspartate (PALA). UDP is then converted
into either thymidine monophosphate (TMP) or cytidine triphosphate (CTP), the latter via uridine triphospate (UTP). De novo
TMP synthesis requires DHFR, so the reaction can be blocked in the same way as in de novo purine synthesis, making cell survival
dependent on the salvage enzyme thymidine kinase (TK). Thus, the Tk gene can be used as a selectable marker. There is no salvage
pathway for CTP.
can only be used with mutant cell lines in which the
corresponding host gene is non-functional. This
restricts the range of cells that can be transfected.
Endogenous markers have therefore been largely
superseded by so-called dominant selectable markers,
which confer a phenotype that is entirely novel
to the cell and hence can be used in any cell type.
Such markers are usually drug-resistance genes of
bacterial origin, and transformed cells are selected on
a medium that contains the drug at an appropriate
concentration. For example, Escherichia coli transposons Tn5 and Tn601 contain distinct genes encoding neomycin phosphotransferase, whose expression
confers resistance to aminoglycoside antibiotics
(kanamycin, neomycin and G418). These are proteinsynthesis inhibitors, active against bacterial and
eukaryotic cells, and can therefore be used for selection in either bacteria or animals. By attaching the
POGC10 9/11/2001 11:04 AM Page 179
Gene transfer to animal cells
179
Table 10.1 Commonly used endogenous selectable marker genes. Most of these markers are involved in the redundant
endogenous nucleotide biosynthetic pathways (Fig. 10.1). They can also be used as counterselectable markers. For
example, negative selection for TK activity is achieved using toxic thymidine analogs (e.g. 5-bromodeoxyuridine,
ganciclovir), which are incorporated into DNA only if there is TK activity in the cell.
Marker
Product
Selection
References
Ada
Adenosine deaminase
Xyl-A (9-b-d-xylofuranosyl adenosine)
and 2′-deoxycoformycin
Kaufman et al. 1986
Aprt
Adenine
phosphoribosyltransferase
Adenine plus azaserine, to block de novo dATP
synthesis
Lester et al. 1980
Cad
Multifunctional enzyme
PALA (N-phosphonacetyl-l-aspartate) inhibits the
aspartate transcarbamylase activity of CAD*
De Saint-Vincent et al.
1981
Hprt
Hypoxanthine-guanine
phosphoribosyltransferase
Hypoxanthine and aminopterin, to block de novo
IMP synthesis
Selected on HAT medium
Lester et al. 1980
Tk
Thymidine kinase
Thymidine and aminopterin to block de novo
dTTP synthesis
Selected on HAT medium
Wigler et al. 1977
*CAD: carbamyl phosphate synthetase/aspartate transcarbamylase/dihydroorotase.
selectable marker to the simian virus 40 (SV40)
early promoter (Berg 1981) or the HSV Tk promoter
(Colbère-Garapin et al. 1981), both of which function
in many cell types, neomycin phosphotransferase
was shown to confer antibiotic resistance in a variety of non-mutant cell lines. This marker continues
to be used in many contemporary expression vectors. The power of aminoglycoside antibiotic resistance as a selective system in eukaryotes is now very
evident. It also has application in yeast (Chapter 9)
and plants (Chapter 12). Other commonly used
dominant selectable markers are listed in Table 10.2.
Selectable markers and transgene amplification
If animal cells are exposed to toxic concentrations
of certain drugs, rare individual cells can survive
because they have spontaneously undergone a
mutation that confers resistance to that drug. The
first such compound to be investigated in this manner was the folic acid analog methotrexate, which
is a competitive inhibitor of the enzyme dihydrofolate reductase (DHFR). The analysis of surviving cells
showed that some had undergone point mutations
in the Dhfr locus, producing an enzyme with resistance to the inhibitor. Others had undergone muta-
tions in other loci, for example preventing the
uptake of the drug. The most interesting group comprised those cells that had survived by amplifying
the Dhfr locus, therefore providing enough enzyme
to outcompete the inhibitor (Schimke et al. 1978).
This type of amplification mutation is useful because
the drug dose can be progressively increased, resulting in the stepwise selection of cells with massively
amplified Dhfr gene arrays. Such cells can survive in
media containing methotrexate concentrations up
to 10 000 times higher than the nominal dose lethal
to wild-type cells. The amplified loci are often maintained within the chromosome as extended homogeneously staining regions or alternatively as small
extra chromosomes called double minutes.
Cells with high copy numbers of the Dhfr locus are
not generated in response to methotrexate exposure,
but arise randomly in the population and are selected
on the basis of their resistance to the drug. The
random nature of the amplification is confirmed by
the fact that, as well as the Dhfr gene itself, extensive
regions of flanking DNA are also amplified, even
though they confer no advantage on the cell.
This phenomenon can be exploited to co-amplify
transgenes introduced along with a Dhfr marker
gene, resulting in high-level expression. Wigler et al.
POGC10 9/11/2001 11:04 AM Page 180
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CHAPTER 10
Table 10.2 Commonly used dominant selectable marker genes in animals (see Box 12.2 for selectable markers
used in plants).
Marker
Product (and source)
Principles of selection
References
as
Asparagine synthase
(Escherichia coli )
Toxic glutamine analog albizziin
Andrulis &
Siminovitch 1981
ble
Glycopeptide-binding protein
(Streptoalloteichus hindustantus)
Confers resistance to glycopeptide antibiotics
bleomycin, pheomycin, ZeocinTM
Genilloud et al. 1984
bsd
Blasticidin deaminase
(Aspergillus terreus)
Confers resistance to basticidin S
Izumi et al. 1991
gpt
Guanine-xanthine
phosphoribosyltransferase
(E. coli )
Analogous to Hprt in mammals, but possesses
additional xanthine phosphoribosyltransferase
activity, allowing survival in medium containing
aminopterin and mycophenolic acid
(Fig 10.1)
Mulligan & Berg
1981b
hisD
Histidinol dehydrogenase
(Salmonella typhimurium)
Confers resistance to histidinol
Mantei et al. 1979
hpt
Hygromycin phosphotransferase
(E. coli )
Confers resistance to hygromycin-B
Blochlinger &
Diggelmann 1984
neo (nptII)
Neomycin phosphotransferase
(E. coli )
Confers resistance to aminoglycoside
antibiotics (e.g. neomycin, kanamycin, G418)
Colbère-Garapin et al.
1981
pac
Puromycin N-acetyltransferase
(Streptomyces alboniger)
Confers resistance to puromycin
Vara et al. 1986
trpB
Tryptophan synthesis (E. coli )
Confers resistance to indole
Hartman & Mulligan
1988
(1980) demonstrated this principle by transfecting
methotrexate-sensitive mouse cells with genomic
DNA from the methotrexate-resistant cell line A29,
which contains multiple copies of an altered Dhfr
gene. They linearized the A29 genomic DNA with
the restriction enzyme SalI, and ligated it to SalIlinearized pBR322 DNA prior to transfection. Following stepwise drug selection of the transformed
cells, Southern-blot hybridization showed that the
amount of pBR322 DNA had increased more than
50-fold.
Methotrexate selection has been used for the
large-scale expression of many recombinant proteins,
including tissue plasminogen activator (Kaufman
et al. 1985), hepatitis B surface antigen (HBSAg)
(Patzer et al. 1986) and human factor VIII (Kaufman
et al. 1988). CHO cells are preferred as hosts for this
expression system because of the availability of a
number of dhfr− mutants (Urlaub et al. 1983). In
non-mutant cell lines, non-transformed cells can
survive selection by amplifying the endogenous Dhfr
genes, generating a background of ‘false positives’.
Alternative strategies have been developed that
allow DHFR selection to be used in wild-type cells.
Expressing the Dhfr marker using a strong constitutive promoter (Murray et al. 1983) or using a
methotrexate-resistant allele of the mouse gene
(Simonsen & Levinson 1983) allows selection at
methotrexate concentrations much higher than
wild-type cells can tolerate. The E. coli dhfr gene is
also naturally resistant to methotrexate, although
for this reason it cannot be used for amplifiable selection (O’Hare et al. 1981). Another useful strategy is
to employ a second marker gene, such as neo, allowing non-transformed cells to be eliminated using
G418 (Kim & Wold 1985). Although dhfr is the most
widely used amplifiable marker, many others have
been evaluated, as shown in Table 10.3.
POGC10 9/11/2001 11:04 AM Page 181
Gene transfer to animal cells
181
Table 10.3 Common markers used for in situ gene amplification. Many amplifiable markers can also be used as
endogenous or dominant selectable markers, but, in some cases, the drug used for amplification may not be the same as
that used for standard selection.
Product
Marker
Amplifying selective drug
References
Ada
Adenosine deaminase
Deoxycoformycin
Kaufman et al. 1986
as
Asparagine synthase
b-Aspartylhydroxamate
Cartier et al. 1987
Cad
Aspartate transcarbamylase
N-Phosphonacetyl-l-aspartate
Wahl et al. 1984
Dhfr
Dihydrofolate reductase
Methotrexate
Kaufman et al. 1985
gpt
Xanthine-guanine phosphoribosyltransferase
Mycophenolic acid
Chapman et al. 1983
GS
Glutamine synthase
Methionine sulphoxamine
Cockett et al. 1990
Hprt
Hypoxanthine-guanine
phosphoribosyltransferase
Aminopterin
Kanalas & Suttle 1984
Impdh
Inosine monophosphate dehydrogenase
Mycophenolic acid
Collart & Huberman 1987
Mt-1
Metallothionein 1
Cd2+
Beach & Palmiter 1981
Mres
Multidrug resistance: P-glycoprotein 170 gene
Adriamycin, colchicine, others
Kane et al. 1988
Odc
Ornithine decarboxylase
Difluoromethylornithine
Chiang & McConlogue 1988
Tk
Thymidine kinase
Aminopterin
Roberts & Axel 1982
Umps
Uridine monophosphate synthases
Pyrazofurin
Kanalas & Suttle 1984
Plasmid vectors for DNA-mediated gene transfer
Stable transformation by integration can be achieved
using any source of DNA. The early gene-transfer
experiments discussed above were carried out using
complex DNA mixtures, e.g. genomic DNA, bacterial
plasmids and phage. Calcium phosphate transfection was used in most of these experiments, and
the specific donor DNA was often bulked up with a
non-specific carrier, such as cleaved salmon-sperm
DNA. However, it is generally more beneficial to use
a purified source of the donor transgene. This principle was originally demonstrated by Wigler et al.
(1977), who transfected cultured mouse cells with
a homogeneous preparation of the HSV Tk gene.
Later, this gene was cloned in E. coli plasmids to provide a more convenient source. The use of plasmid
vectors for transfection provides numerous other
advantages, depending on the modular elements
included on the plasmid backbone.
1 The convenience of bacterial plasmid vectors can
be extended to animal cells, in terms of the ease of
subcloning, in vitro manipulation and purification of
recombinant proteins (p. 76).
2 More importantly, modular elements can be
included to drive transgene expression, and these
can be used with any transgene of interest. The pSV
and pRSV plasmids are examples of early expression
vectors for use in animal cells. As discussed in
Box 10.2, transcriptional control sequences from
SV40 and Rous sarcoma virus are functional in a
wide range of cell types. The incorporation of
these sequences into pBR322 generated convenient
expression vectors in which any transgene could be
controlled by these promoters when integrated into
the genome of a transfected cell (Fig. 10.2).
3 The inclusion of a selectable marker gene
obviates the need for cotransformation, since the
transgene and marker remain linked when they
cointegrate into the recipient cell’s genome. A range
of pSV and pRSV vectors were developed containing
alternative selectable marker genes, e.g. pSV2-neo
(Southern & Berg 1982), pSV2-gpt (Mulligan & Berg
1980) and pSV2-dhfr (Subramani et al. 1981).
POGC10 9/11/2001 11:04 AM Page 182
182
CHAPTER 10
late
am
r
p
SV
4
0
lyA
po
AAAA
SV40 small
T intron
pSV2-dhfr
(5.0 kb)
4 As discussed below, some plasmid vectors for gene
transfer to animal cells are designed to be shuttle
vectors, i.e. they contain origins of replication
functional in animal cells, allowing the vector to be
maintained as an episomal replicon.
Non-replicating plasmid vectors for
transient transformation
o ri
PE
fr
dh
SV40 ori
late
p
SV
4
0
lyA
po
am
r
smal
SV40 ronl
T int
AAAA
pRSVneo
(5.6 kb)
o ri
PLTR
LTR
5n
Tn
eg
Fig. 10.2 The mammalian plasmid expression vectors
pSV2-dhfr and pRSV-neo.
One application in which the use of plasmid vectors
is critical is transient transformation. Here, the goal
is to exploit the short-term persistence of extrachromosomal DNA. Such experiments have a variety of
uses, including transient assays of gene expression
and the recovery of moderate amounts of recombinant protein. Generally, transient transformation
is used as a test system, e.g. to assay regulatory
elements using reporter genes (Box 10.1), to check
the correct function of an expression construct
before going to the expense of generating stable cell
lines or to recover moderate amounts of recombinant protein for verification purposes. Transient
transformation is particularly useful for testing large
numbers of alternative constructs in parallel. No
regime of selection is required because stable cell
lines are not recovered – the cells are generally
transfected, assayed after 1 or 2 days and then discarded. The simplest way to achieve the transient
transformation of animal cells is to use a plasmid
Box 10.1 Reporter genes and promoter analysis
Reporter genes are also known as screenable marker
genes. These differ from selectable marker genes
in that they do not confer a property that allows
transformed cells to survive under selective
conditions. Instead, they encode a product that
can be detected using a simple and inexpensive
assay.
When controlled by a strong constitutive promoter,
reporter genes are often used as markers to confirm
transient or stable transformation, since only cells
containing the reporter-gene construct can express
the corresponding protein. Importantly, the assays
used to detect reporter-gene activity are quantitative,
so they can also be used to measure transformation
efficiency. If attached to a cloned promoter,
reporter genes can therefore be used to determine
transcriptional activity in different cell types and
under different conditions. Transient reporter assays
have been widely used to characterize and dissect
the regulatory elements driving eukaryotic genes, as
shown in the example below. The use of reporters is
advantageous, because it circumvents the necessity
to derive different assays for individual genes
and also allows the activities of transgenes and
homologous endogenous genes to be distinguished
in the same cell.
continued
POGC10 9/11/2001 11:04 AM Page 183
Gene transfer to animal cells
183
Box 10.1 continued
An example of in vitro promoter analysis using
chloramphenicol acetyltransferase
The first reporter gene to be used in animal cells was
cat, derived from E. coli transposon Tn9 (Gorman
et al. 1982b); it has also been used to a certain
extent in plants (Herrera-Estrella et al. 1983a,b).
This gene encodes the enzyme chloramphenicol
acetyltransferase (CAT), which confers resistance to
the antibiotic chloramphenicol by transferring acetyl
groups on to the chloramphenicol molecule from
acetyl-CoA. If 14C-labelled chloramphenicol is used
as the substrate, CAT activity produces a mixture
of labelled monoacetylated and diacetylated
forms, which can be separated by thin-layer
chromatography and detected by autoradiography.
The higher the CAT activity, the more acetylated
forms of chloramphenicol are produced. These
can be quantified in a scintillation counter or using a
phosphorimager. Gorman et al. (1982a,b) placed the
cat gene downstream of the SV40 and Rous sarcoma
virus (RSV) promoters in the expression vectors pSV2
and pRSV2, to create the pSV-CAT and pRSV-CAT
constructs, respectively. These vectors, and derivatives
thereof, have been widely used to analyse transienttransfection efficiency, because the promoters are
active in many animal cells and CAT activity can
be assayed rapidly in cell homogenates.
The cat gene has also been used to test regulatory
elements by attaching it to a ‘minimal promoter’,
typically a simple TATA box. This basic construct
generates only low-level background transcription
in transiently transfected cells. The activity of
other regulatory elements, such as promoters and
enhancers, and response elements, which activate
transcription in response to external signals, can be
tested by subcloning them upstream of the minimal
promoter and testing their activity in appropriate cell
types. In an early example of this type of experiment,
Walker et al. (1986) attached the promoter and
5′-flanking sequences of human and rat insulin and
rat chymotrypsin genes, which are expressed at a
high level only in the pancreas, to the cat gene. Each
gene is expressed in clearly distinct cell types: insulin
is synthesized in endocrine β cells and chymotrypsin
in exocrine cells. Plasmid DNA was introduced into
either pancreatic endocrine or pancreatic exocrine
cell lines in culture and, after a subsequent 44 h
incubation, cell extracts were assayed for CAT
activity. It was found that the constructs retained
their preferential expression in the appropriate cell
type. The insulin 5′-flanking DNA conferred a high
level of CAT expression in the endocrine but not the
exocrine cell line, with the converse being the case for
the chymotrypsin 5′-flanking DNA. The analysis was
extended by creating deletions in the 5′-flanking
sequences and testing their effects on expression.
From such experiments it could be concluded that
sequences located 150–300 bp upstream of the
transcription start are essential for appropriate
cell-specific transcription.
Other reporter genes
The cat reporter gene has been widely used for in
vitro assays but has a generally low sensitivity and is
dependent on a rather cumbersome isotopic assay
format. An alternative reporter gene, SeAP (secreted
alkaline phosphatase), has been useful in many cases
because various sensitive colourimetric, fluorometric
and chemiluminescent assay formats are available.
Also, since the reporter protein is secreted, it can be
assayed in the growth medium, so there is no need
to kill the cells (Berger et al. 1988, Cullen & Malim
1992). The bacterial genes lacZ and gusA have
been used as reporters for in vitro assays using
colourimetric and fluorometric substrates. An
important advantage of these markers is that they
can also be used for in situ assays, since histological
assay formats are available. More recently,
bioluminescent markers, such as luciferase and green
fluorescent protein, have become popular because
these can be assayed in live cells and whole animals
and plants. The lacZ, gusA, luciferase and green
fluorescent protein reporter genes are discussed in
more detail in Box 13.1.
POGC10 9/11/2001 11:04 AM Page 184
CHAPTER 10
rip
tra
ns
c
vp
2
3’
A
vp
3
NA
18 S m
RN
A
19S mR
NA
A
A
AA
3’
A
AA
A
Transient transformation can also be achieved using
replicon vectors that contain origins of replication
derived from certain viruses of the polyomavirus
family, such as SV40 and the murine polyomavirus. These viruses cause lytic infections, i.e. the
viral genome replicates to a very high copy number,
resulting in cell lysis and the release of thousands of
progeny virions. During the infection cycle, viral
gene products accumulate at high levels, so there
has been considerable interest in exploiting this
strategy to produce recombinant proteins. SV40
was the first animal virus to be characterized in
detail at the molecular level, and for this reason it
was also the first to be developed as a vector. The
productive host range of the virus is limited to certain simian cells. The development of SV40 replicon
vectors is discussed in some detail below, but first it is
necessary to understand a little of the molecular
biology of the virus itself.
SV40 has a small icosahedral capsid and a circular double-stranded DNA genome of approximately
5 kb. The genome has two transcription units, known
as the early and late regions, which face in opposite
directions. Both transcripts produce multiple products by alternative splicing (Fig. 10.3). The early
region produces regulatory proteins, while the late
region produces components of the viral capsid.
vp
1
AA
Runaway polyomavirus replicons
ts
AA
Transformation with replicon vectors
16S mR
AA
vector lacking an origin of replication functional in
the host. Although the vector cannot replicate, gene
expression from a mammalian transcription unit is
possible for as long as the plasmid remains stable,
which depends on the host cell’s propensity to
break down extrachromosomal DNA. Linear DNA
is degraded very quickly in mammalian cells, so
high-quality supercoiled plasmid vectors are used.
Even covalently closed circular DNA tends to remain
stable for only 1 or 2 days in most animal cells, but
this is sufficient for the various transient-expression
assays. Some cell types, however, are renowned for
their ability to maintain exogenous DNA for longer
periods. In the human embryonic kidney cell line
293, for example, supercoiled plasmid DNA can
remain stable for up to 80 h (Gorman et al. 1990).
Late
184
5’
5’
t
T
Origin of
replication
Early transc
s
ript
Fig. 10.3 Transcripts and transcript processing of SV40.
Intron sequences which are spliced out of the transcripts are
shown by red lines.
Transcription is controlled by a complex regulatory
element located between the early and late regions,
and this includes early and late promoters, an
enhancer and the origin of replication. During the
first stage of the SV40 infection cycle, the early transcript produces two proteins, known as the large T
and small t tumour antigens. The function of the T
antigen is particularly important, as this protein
binds to the viral origin of replication and is absolutely required for genome replication. All vectors
based on SV40 must therefore be supplied with
functional T antigen, or they cannot replicate, even
in permissive cells. The T antigen also acts as an
oncoprotein, interacting with the host’s cell-cycle
machinery and causing uncontrolled cell proliferation.
The first SV40 vectors were viral vectors and were
used to introduce foreign genes into animal cells by
transduction. The small size of the viral genome
made in vitro manipulation straightforward. Either
the late region (Goff & Berg 1976) or the early region
(Gething & Sambrook 1981) could be replaced with
foreign DNA. However, since both these regions are
essential for the infection cycle, their functions had
to be provided in trans initially by a co-introduced
helper virus. The use of early replacement vectors
was considerably simplified by the development of
POGC10 9/11/2001 11:04 AM Page 185
T7
SV4
0
V
P CM
i nt
ro
n/
Ampicilli
n
pA
Poly
oma ori
pcDNA1.1/
Amp
4.8 kb
3
M1
i
or
the COS cell line, a derivative of the African greenmonkey cell line CV-1 containing an integrated
partial copy of the SV40 genome. The integrated
fragment included the entire T-antigen coding
sequence and provided this protein in trans to any
SV40 recombinant in which the early region had
been replaced with foreign DNA (Gluzman 1981).
For example, using this system, Gething and Sambrook (1981) made recombinant viruses that expressed influenza-virus haemagglutinin in COS cells.
The major problem with these initial SV40 vectors was that the capacity of the viral capsid allowed
a maximum of only about 2.5 kb of foreign DNA to
be incorporated. The discovery that plasmids carrying the SV40 origin of replication behaved in the
same manner as the virus itself, i.e. replicating to a
high copy number in permissive monkey cells, was
a significant breakthrough. Since these SV40 replicons were not packaged into viral capsids, there
were no size constraints on the foreign DNA. Many
laboratories developed plasmid vectors on this
principle (Myers & Tjian 1980; Lusky & Botchan
1981; Mellon et al. 1981). In general, these vectors
consisted of a small SV40 DNA fragment (containing the origin of replication) cloned in an E. coli plasmid vector. Some vectors also contained a T-antigen
coding region and could be used in any permissive
cell line, while others contained the origin alone and
could only replicate in COS cells. Permanent cell
lines are not established when SV40 replicons are
transfected into COS cells, because the massive
vector replication eventually causes cell death. However, even though only a low proportion of cells are
transfected, the high copy number (105 genomes
per cell) is compensatory, allowing the transient
expression of cloned genes and the harvesting of
large amounts of recombinant protein.
Similar vectors have been designed that incorporate the murine-polyomavirus origin, which is functional in mouse cells. These vectors allow transient,
high-level recombinant-protein expression in permissive mouse cells, such as MOP-8 (which contains
an integrated polyomavirus genome and supplies
T antigen in trans (Muller et al. 1984)) and WOP
(which is latently infected with the virus). A series
of vectors currently marketed by Invitrogen are
particularly versatile, because they contain both
the SV40 and the murine-polyomavirus origins,
185
Hind III
BamH I
BstX I
EcoR I
EcoR V
BstX I
Not I
Xho I
Sph I
Nsi I
Xba I
Gene transfer to animal cells
ColE1
ri
0o
SV 4
Fig. 10.4 The transient expression vector pcDNA1.1/Amp,
which has origins of replication for both SV40 and murine
polyomavirus. (Reproduced with permission from
Invitrogen.)
allowing high-level protein expression in cells that
are permissive for either virus. For example, the
vector pcDNA1.1/Amp contains the SV40 and
polyoma origins, a transcription unit comprising the
human cytomegalovirus promoter and SV40 intron/
polyadenylaton site, an interstitial polylinker to insert
the transgene and the ampicillin-resistance marker
for selection in E. coli (Fig. 10.4).
Stable transformation with BK and BPV replicons
As an alternative to integration, stable transformation can be achieved using a recombinant vector
that is maintained as an episomal replicon. While
viruses such as SV40 use a lytic infection strategy,
others, such as human BK virus and Epstein–Barr
virus (EBV), cause latent infections, where the viral
genome is maintained as a low- to moderatecopy-number replicon that does not interfere with
host cell growth. Plasmids that contain such latent
origins behave in a similar manner to the parental
virus, except they are not packaged in a viral capsid.
Such vectors are advantageous, because the DNA
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does not need to integrate in order to be stably
maintained, thus stable transformation occurs with
an efficiency equal to that normally achieved with
transient transformation. Furthermore, while the
expression of integrated transgenes is often affected
by the surrounding DNA, a problem known as the
position effect (see Box 11.1), these episomal vectors
are not subject to such influences.
The human BK polyomavirus infects many cell
types and is maintained with a copy number of about
500 genomes per cell. Plasmid vectors containing
the BK origin replicate in the same manner as the
virus when the BK T antigen is provided in trans.
However, by incorporating a selectable marker such
as neo into the vector, it is possible to progressively
increase the concentration of antibiotic in the medium
and select for cells with higher vector copy number.
Cell lines have been propagated for over a year under
such conditions (De Benedetti & Rhoads 1991) and
stable lines with up to 9000 copies of the genome
have been produced. Any transgene incorporated
into the vector is expressed at very high levels, leading to the recovery of large amounts of recombinant
protein (Sabbioni et al. 1995). SV40 itself can also
be used as a stable episomal vector if the availability
of T antigen is rationed to prevent runaway replication. This has been achieved using a conditional
promoter and by mutating the coding region to
render the protein temperature-sensitive (Gerard &
Gluzman 1985, Rio et al. 1985).
The first virus to be developed as an episomal
replicon was bovine papillomavirus (BPV). The
papillomaviruses are distantly related to the polyomaviruses and cause papillomas (warts) in a range
of mammals. BPV has been exploited as a stable
expression vector because it can infect mouse cells
without yielding progeny virions. Instead, the viral
genome is maintained as an episomal replicon, with
a copy number of about 100. The molecular biology
of BPV is considerably more complex that that of
SV40, but the early part of the infection cycle is similar, involving the production of a T antigen that:
(i) is required for viral replication; and (ii) causes
oncogenic transformation of the host cell. The early
functions of BPV are carried on a 5.5 kb section of
the genome, which is called the 69% transforming
fragment (BPV69T). This was cloned in the E. coli
plasmid pBR322 and was shown to be sufficient to
establish and maintain episomal replication, as well
as inducing cell proliferation (Sarver et al. 1981a).
Initially, the ability of the virus to cause uncontrolled cell proliferation was used to identify transformed cells, but this limited the range of cell types
that could be used. The incorporation of selectable
markers, such as neo, allows transformants to be
selected for resistance to G418, permitting the use of
a wider range of cell types (Law et al. 1983). BPV69T
replicons have been used to express numerous proteins, including rat preproinsulin (Sarver et al. 1981b)
and human β-globin (Di Maio et al. 1982). Generally, the plasmids are maintained episomally, but
the copy number varies from 10 to over 200 vector
molecules per cell. Some investigators have reported
the long-term maintenance of such episomal transformants (Fukunaga et al. 1984), while others found
a tendency for the construct to integrate into the
genome (Ostrowski et al. 1983, Sambrook et al.
1985). Recombination within the vector or between
vectors is also a fairly common observation, resulting
in unpredictable spontaneous deletions and plasmid
oligomerization. Recombination events and copy
number appear to be affected by multiple factors,
including the host cell type, the incorporated transgene and the structure of the vector itself (see Mecsas
& Sugden 1987).
Stable transformation with EBV replicons
Unlike BPV replicons, vectors based on EBV replicate
very stably in mammalian cells. EBV is a herpesvirus
(see also discussion of herpes simplex virus later in
the chapter), with a large double-stranded DNA
genome (approximately 170 kb), which predominantly infects primate and canine cells. It is also
naturally lymphotrophic, infecting B cells in humans
and causing infectious mononucleosis. In cultured
lymphocytes, the virus becomes established as an
episomal replicon with about 1000 copies per cell
(Miller 1985). Although the virus itself only infects
lymphocytes, the genome is maintained in a wide
range of primate cells if introduced by transfection.
Only two relatively small regions of the genome are
required for episomal maintenance – the latent
origin (oriP) and a gene encoding a trans-acting
regulator called Epstein–Barr nuclear antigen 1
(EBNA1) (Yates et al. 1984, Reisman et al. 1985).
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Gene transfer to animal cells
These sequences have formed the basis of a series of
latent EBV-based plasmid expression vectors, which
are maintained at a copy number of two to 50 copies
per cell. The first EBV vectors comprised the oriP
element cloned in a bacterial plasmid and could
replicate only if EBNA1 was supplied in trans. Yates
et al. (1984) described the construction of a shuttle
vector, pBamC, comprising oriP, a bacterial origin
and ampicillin-resistance gene derived from pBR322,
and the neomycin phosphotransferase marker for
selection in animal cells. A derivative vector, pHEBO,
contained a hygromycin-resistance marker, instead
of neo (Sugden et al. 1985). The presence of a
selectable marker in such vectors is important,
because, if selection pressure is not applied, EBV
replicons are lost passively from the cell population
at about 5% per cell generation. Yates et al. (1985)
added the EBNA1 gene to pHEBO, producing a construct called p201 that was capable of replicating
independently. Similar constructs have been developed in other laboratories (Lupton & Levine 1985).
EBV replicons have been used to express a wide
range of proteins in mammalian cell lines, including
the epidermal growth-factor receptor (Young et al.
1988), the tumour-necrosis-factor receptor (Heller
et al. 1990) and an Na+K+ ATPase (Canfield et al.
1990). Generally, such studies have resulted in highlevel and long-term gene expression (reviewed by
Margolskee 1992, Sclimenti & Calos 1998). It has
been suggested that the stability of EBV replicons
may reflect the fact that replication is limited to once
per cell cycle, unlike BPV replicons, which replicate
continuously throughout the S phase (Gilbert &
Cohen 1987, DuBridge & Calos 1988). This, together
with the large genome size, allows EBV-derived
vectors to carry large DNA fragments, including
mammalian complementary DNAs (cDNAs) and
genes. EBV replicons have therefore been used
for the construction of episomal cDNA libraries
for expression cloning (Margolskee et al. 1988)
and, more recently, for the preparation of genomic
libraries (Sun et al. 1994). The oriP element has also
been incorporated into YAC vectors carrying large
human genomic DNA fragments. These linear
vectors were then circularized in vitro and introduced into human cells expressing EBNA1, whereupon they were maintained as episomal replicons
(Simpson et al. 1996). EBV can also replicate lytic-
187
ally in primate B cells, and this requires a separate
origin (oriLyt) and a distinct trans-acting regulator
called ZEBRA. Vectors have been developed containing both origins, and these are maintained at a moderate copy number in cells latently infected with the
virus, but can be amplified up to 400-fold if ZEBRA
is supplied by transfecting the cells with the corresponding viral gene under the control of a constitutive promoter (Hammerschmidt & Sugden 1988).
Interestingly, while the latent vector remains as a
circular replicon, vector DNA isolated from induced
cells is present as multicopy concatemers, reflecting
the rolling-circle mechanism of lytic replication.
Gene transfer by viral transduction
General principles of viral vectors
This section describes the use of viruses as vectors for
transduction, i.e. the introduction of genes into animal cells by exploiting the natural ability of the virus
particle, within which the transgene is packaged, to
adsorb to the surface of the cell and gain entry. Due
to the efficiency with which viruses can deliver their
nucleic acid into cells and the high levels of replication and gene expression it is possible to achieve,
viruses have been used as vectors not only for gene
expression in cultured cells but also for gene transfer
to living animals. Four classes of viral vector have
been developed for use in human gene therapy
and have reached phase 1 clinical trials. These are
the retrovirus, adenovirus, herpesvirus and adenoassociated virus (AAV) vectors (Robbins et al. 1998).
Before introducing the individual vector systems,
we discuss some general properties of viral transduction vectors. Transgenes may be incorporated into
viral vectors either by addition to the whole genome
or by replacing one or more viral genes. This is
generally achieved either by ligation (many viruses
have been modified to incorporate unique restriction
sites) or homologous recombination. If the transgene is added to the genome or if it replaces one or
more genes that are non-essential for the infection
cycle in the expression host being used, the vector is
described as replication-competent or helper-independent,
because it can propagate independently. However, if the transgene replaces an essential viral
gene, this renders the vector replication-defective or
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helper-dependent, so that missing functions must be
supplied in trans. This can be accomplished by cointroducing a helper virus or transfecting the cells
with a helper plasmid, each of which must carry the
missing genes. Usually steps are taken to prevent the
helper virus completing its own infection cycle, so
that only the recombinant vector is packaged. It is
also desirable to try and prevent recombination occurring between the helper and the vector, as this can
generate wild-type replication-competent viruses as
contaminants. An alternative to the co-introduction
of helpers is to use a complementary cell line, which
is transformed with the appropriate genes. These
are sometimes termed ‘packaging lines’. For many
applications, it is favourable to use vectors from
which all viral coding sequences have been deleted.
These amplicons (also described as ‘gutless vectors’)
contain just the cis-acting elements required for
packaging and genome replication. The advantage
of such vectors is their high capacity for foreign DNA
and the fact that, since no viral gene products are
made, the vector has no intrinsic cytotoxic effects.
The choice of vector depends on the particular
properties of the virus and the intended host,
whether transient or stable expression is required
and how much DNA needs to be packaged. For
example, icosahedral viruses such as adenoviruses
and retroviruses package their genomes into preformed capsids, whose volume defines the maximum
amount of foreign DNA that can be accommodated.
Conversely, rod-shaped viruses such as the baculoviruses form the capsid around the genome, so there
are no such size constraints. There is no ideal virus
for gene transfer – each has its own advantages and
disadvantages. However, in recent years, a number
of hybrid viral vectors have been developed incorporating the beneficial features of two or more viruses.
The interested reader can consult recent reviews on
this subject (Robbins et al. 1998, Reynolds et al.
1999). Also note that most of the principles discussed
above also apply to the use of plant viruses as vectors
(Chapter 12).
Adenovirus
Adenoviruses are DNA viruses with a linear, doublestranded genome of approximately 36 kb. The genome
of serotype Ad5, from which many adenovirus
vectors are derived, is shown in Fig. 10.5. There are
six early-transcription units, most of which are
essential for viral replication, and a major late
transcript that encodes components of the capsid.
Adenoviruses have been widely used as gene transfer and expression vectors, because they have many
advantageous features, including stability, a high
capacity for foreign DNA, a wide host range that
includes non-dividing cells, and the ability to produce high-titre stocks (up to 1011 plaque-forming
units (pfu)/ml) (reviewed by Berkner 1992). They
are suitable for transient expression in dividing cells
because they do not integrate efficiently into the
genome, but prolonged expression can be achieved
in post-mitotic cells, such as neurons (e.g. see
Davidson et al. 1993, LaSalle et al. 1993). Adenoviruses are particularly attractive as gene therapy
vectors, because the virions are taken up efficiently
by cells in vivo and adenovirus-derived vaccines have
been used in humans with no reported side-effects.
However, the recent death of a patient following
an extreme inflammatory response to adenoviral
TL
L2
L1
E1a
3’
5’
0
Ψ
MLT
E1b
VA
L3
L4
plX
L5
E3
5’
3’
10
20
IVa2
30
40
50
60
70
80
E2a
E2b
90
100
E4
Fig. 10.5 Map of the adenovirus genome, showing the positions of the early transcription units (E), the major late transcript
(MLT), the tripartite leader (TL) and other genes (VA, pIX, IVa2). Terminal repeats are shown in pink, ψ is the packaging site.
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Gene transfer to animal cells
gene-therapy treatment underlines the necessity for
rigorous safety testing (Marshall 1999). A number
of strategies are being developed to control the activity of the immune system when the vectors are first
introduced into the host (reviewed by Benihoud et al.
1999).
Most early adenoviral vectors were replicationdeficient, lacking the essential E1a and E1b genes
and often the non-essential gene E3. These firstgeneration ‘E1 replacement vectors’ had a maximum capacity of about 7 kb and were propagated
in the human embryonic kidney line 293. This is
transformed with the leftmost 11% of the adenoviral
genome, comprising the E1 transcription unit, and
hence supplies these functions in trans (Graham et al.
1977). Although these vectors have been used
with great success, they suffer from two particular
problems: cytotoxic effects, resulting from low-level
expression of the viral gene products, and the
tendency for recombination to occur between the
vector and the integrated portion of the genome,
resulting in the recovery of replication-competent
viruses. Higher-capacity vectors have been developed, which lack the E2 or E4 regions in addition to
E1 and E3, providing a maximum cloning capacity
of about 10 kb. These must be propagated on complementary cell lines providing multiple functions,
and such cell lines have been developed in several
laboratories (e.g. Brough et al. 1996, Gao et al.
1996, Gorziglia et al. 1996, Zhou et al. 1996). The
use of E1/E4 deletions is particularly attractive, as
the E4 gene is responsible for many of the immunological effects of the virus (Gao et al. 1996, Dedieu
et al. 1997). Unwanted recombination has been
addressed through the use of a refined complementary cell line transformed with a specific DNA fragment corresponding exactly to the E1 genes (Imler
et al. 1996). An alternative strategy is to insert
a large fragment of ‘stuffer DNA’ into the nonessential E3 gene, so that recombination yields a
genome too large to be packaged (Bett et al. 1993).
Gutless adenoviral vectors are favoured for in vivo
gene transfer, because they have a large capacity
(up to 37 kb) and minimal cytotoxic effects (reviewed
by Morsey & Caskey 1999). Therefore, transgene
expression persists for longer than can be achieved
using first-generation vectors (e.g. see Schneider
et al. 1998). Complementary cell lines supplying all
189
adenoviral functions are not available at present, so
gutless vectors must be packaged in the presence of a
helper virus, which presents a risk of contamination.
Adeno-associated virus
AAV is not related to adenovirus, but is so called
because it was first discovered as a contaminant in
an adenoviral isolate (Atchison et al. 1965). AAV
is a single-stranded DNA virus, a member of the
parvovirus family, and is naturally replicationdefective, such that it requires the presence of
another virus (usually adenovirus or herpesvirus)
to complete its infection cycle. In adenovirus- or
herpesvirus-infected cells, AAV replicates lytically
and produces thousands of progeny virions (Buller
et al. 1981). However, in the absence of these
helpers, the AAV DNA integrates into the host cell’s
genome, where it remains as a latent provirus (Berns
et al. 1975). In human cells, the provirus integrates
predominantly into the same genetic locus on chromosome 19 (Kotin et al. 1990). Subsequent infection by adenovirus or herpesvirus can ‘rescue’ the
provirus and induce lytic infection.
The dependence of AAV on a heterologous helper
virus provides an unusual degree of control over
vector replication, making AAV theoretically one of
the safest vectors to use for gene therapy. Proviral
integration is considered advantageous for increasing the persistence of transgene expression, while at
the same time the site specificity of this process theoretically limits the chances of insertional mutagenesis. Other advantages include the wide host range,
which encompasses non-dividing cells (reviewed by
Muzyczka 1992, Rabinowitz & Samulski 1998).
The AAV genome is small (about 5 kb) and
comprises a central region containing rep (replicase)
and cap (capsid) genes flanked by 145-base inverted
terminal repeats (Fig. 10.6). In the first AAV vectors, foreign DNA replaced the cap region and
was expressed from an endogenous AAV promoter
(Hermonat & Muzyczka 1984). Heterologous promoters were also used, although in many cases
transgene expression was inefficient because the
Rep protein inhibited their activity (reviewed by
Muzyczka, 1992). Rep interference with endogenous
promoters is also responsible for many of the cytotoxic effects of the virus. Several groups therefore
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p5
p19
p40
rep
polyA
cap
ori, res, pac, int
Fig. 10.6 Map of the adeno-associated virus genome,
showing the three promoters, the rep and cap genes, and the
cis-acting sites required for replication (ori), excision (rescue),
(res), packaging ( pac) and integration (int). Terminal repeats
are shown as pink blocks.
developed vectors in which both genes were deleted
and the transgene was expressed from either an endogenous or a heterologous promoter (McLaughlin
et al. 1988, Samulski et al. 1989). From such experiments, it was demonstrated that the repeats are the
only elements required for replication, transcription,
proviral integration and rescue. All current AAV
vectors are based on this principle. In vitro manipulation of AAV is facilitated by cloning the inverted
terminal repeats in a plasmid vector and inserting
the transgene between them. Traditionally, recombinant viral stocks are produced by transfecting this
construct into cells along with a helper plasmid to
supply AAV products, and then infecting the cells
with adenovirus to stimulate lytic replication and
packaging. This has generally yielded recombinant
AAV titres too low to use for human gene therapy
and contaminated with helper AAV and adenovirus.
The recent development of transfection-based adenoviral helper plasmids, packaging lines and the use
of affinity chromatography to isolate AAV virions
has helped to alleviate such problems (reviewed by
Monahan & Samulski 2000).
AAV vectors have been used to introduce genes
efficiently into many cell types, including liver (Snyder
et al. 1997), muscle (Pruchnic et al. 2000) and
neurons (Davidson et al. 2000). However, deletion
of the rep region abolishes the site specificity of proviral integration, so the vector integrates at essentially
random positions, which may increase the risk of
insertional gene inactivation (Weitzman et al. 1994,
Yang et al. 1997, Young et al. 2000). It is also
unclear whether the persistence of the vector and
prolonged transgene expression are due primarily
to vector integration or to episomal maintenance
of concatemeric double-stranded DNA (dsDNA)
copies of the genome (for discussion see Monahan &
Samulski 2000). The fact that AAV uses concatemeric replication intermediates has been used to
circumvent perhaps the most serious disadvantage
of AAV vectors, which is the limited capacity for
foreign DNA. This strategy involves cloning a large
cDNA as two segments in two separate vectors,
which are co-introduced into the same cell. The 5′
portion of the cDNA is cloned in one vector, downstream of a promoter and upstream of a splice donor
site. The 3′ portion is cloned in another vector,
downstream of a splice acceptor. Concatemerization
results in the formation of heterodimers and transcription across the junction yields an mRNA that
can be processed to splice out the terminal repeats of
the vector. In this way, cDNAs of up to 10 kb can be
expressed (Nakai et al. 2000, Sun et al. 2000).
Baculovirus
Baculoviruses have rod-shaped capsids and large,
dsDNA genomes. They productively infect arthropods, particularly insects. One group of baculoviruses,
known as the nuclear polyhedrosis viruses, have an
unusual infection cycle that involves the production
of nuclear occlusion bodies. These are proteinaceous
particles in which the virions are embedded, allowing the virus to survive harsh environmental conditions, such as desiccation (reviewed by Fraser 1992).
Baculovirus vectors are used mainly for high-level
transient protein expression in insects and insect
cells (King & Possee 1992, O’Reilley et al. 1992). The
occlusion bodies are relevant to vector development
because they consist predominantly of a single
protein, called polyhedrin, which is expressed at
very high levels. The nuclear-occlusion stage of the
infection cycle is non-essential for the productive
infection of cell lines; thus the polyhedrin gene
can be replaced with foreign DNA, which can be
expressed at high levels under the control of the
endogenous polyhedrin promoter. Two baculoviruses have been extensively developed as vectors,
namely the Autographa californica multiple nuclear
polyhedrosis virus (AcMNPV) and the Bombyx mori
nuclear polyhedrosis virus (BmNPV). The former is
used for protein expression in insect cell lines, particularly those derived from Spodoptera frugiperda (e.g.
Sf9, Sf21). The latter infects the silkworm and has
been used for the production of recombinant protein
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Gene transfer to animal cells
in live silkworm larvae. One limitation of this expression system is that the glycosylation pathway in
insects differs from that in mammals, so recombinant mammalian proteins may be incorrectly glycosylated and hence immunogenic (reviewed by Fraser
1992). This has been addressed by using insect cell
lines chosen specifically for their ability to carry out
mammalian-type post-translational modifications,
e.g. those derived from Estigmene acrea (Ogonah et al.
1996). An innovative approach to this problem is to
exploit the indefinite capacity of baculovirus vectors
to coexpress multiple transgenes and thus modify
the glycosylation process in the host cell line by
expressing appropriate glycosylation enzymes along
with the transgene of interest. Wagner et al. (1996)
used this strategy to coexpress fowl-plague haemagglutinin and β-1,2-N-acetylglucosaminyltransferase, resulting in the synthesis of large amounts
of haemagglutinin correctly modified with Nacetylglucosamine residues. Hollister et al. (1998)
have developed an Sf9 cell line that expresses 1,4galactosyltransferase under the control of a baculovirus immediate early promoter, such that gene
expression is induced by baculovirus infection. These
cells were infected with a recombinant baculovirus
vector carrying a tissue plasminogen-activator transgene, resulting in the production of recombinant
protein that was correctly galactosylated.
Polyhedrin gene-replacement vectors are the
most popular due to the high level of recombinant
protein that can be expressed (up to 1 mg/106 cells).
The polyhedrin upstream promoter and 5′ untranslated region are important for high-level foreigngene
expression and these are included in all polyhedrin
replacement vectors (Miller et al. 1983, Maeda et al.
1985). The highest levels of recombinant-protein
expression were initially achieved if the transgene
was expressed as a fusion, incorporating at least
the first 30 amino acids of the polyhedrin protein
(Lucklow & Summers 1988). However, this was
shown to be due not to the stabilizing effects of the
leader sequence on the recombinant protein, but to
the presence of regulatory elements that overlapped
the translational start site. Mutation of the polyhedrin start codon has allowed these sequences to be
incorporated as part of the 5′ untranslated region of
the foreign gene cassette, so that native proteins can
be expressed (Landford 1988). Replacement of the
191
polyhedrin gene also provides a convenient method
to detect recombinant viruses. The occlusion bodies
produced by wild-type viruses cause the microscopic
viral plaques to appear opalescent if viewed under
an oblique light source (OB+), while recombinant
plaques appear clear (OB−) (Smith et al. 1983).
Insertion of the E. coli lacZ gene in frame into the
polyhedrin coding region allows blue–white screening of recombinants in addition to the OB assay
(Pennock et al. 1984), and many current baculovirus expression systems employ lacZ as a screenable
marker to identify recombinants. Substitution of
lacZ with the gene for green fluorescent protein
allows the rapid identification of recombinants
by exposing the plaques to UV light (Wilson et al.
1997).
The construction of baculovirus expression vectors involves inserting the transgene downstream of
the polyhedrin promoter. Since the genome is large,
this is usually achieved by homologous recombination using a plasmid vector carrying a baculovirus
homology region. A problem with homology-based
strategies for introducing foreign DNA into large
viral genomes is that recombinants are generated at
a low efficiency. For baculovirus, recombinant vectors are recovered at a frequency of 0.5–5% of total
virus produced. The proportion of recombinants
has been increased by using linear derivatives of
the wild-type baculovirus genome containing large
deletions, which can be repaired only by homologous recombination with the targeting vector.
Compatible targeting vectors span the deletion
and provide enough flanking homologous DNA to
sponsor recombination between the two elements
and to generate a viable, recombinant genome.
Such approaches result in the production of up to
90% recombinant plaques (Kitts & Possee 1993;
Fig. 10.7). Recently, alternative systems have been
described in which the baculovirus genome is
maintained as a low-copy-number replicon in bacteria or yeast, allowing the powerful genetics of these
microbial systems to be exploited. In the bacterial
system marketed by Gibco-BRL under the name
‘Bac-to-Bac’, the baculovirus genome is engineered
to contain an origin of replication from the E. coli F
plasmid. This hybrid replicon, called a bacmid, also
contains the target site for the transposon Tn7,
inserted in frame within the lacZ gene, which is itself
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CHAPTER 10
130 kb
Wild type AcMNPV DNA
(replication competent)
Deleted polyhedrin gene + flanking
regions (replication defective).
Transfer vector – insert foreign
gene in polyhedrin sequence.
Transfect with derivative AcMNPV DNA
Foreign gene
Polyhedr
in
Homologous recombinant
Fig. 10.7 Procedure for the generation of recombinant baculovirus vectors.
Recombinant AcMNPV DNA
(replication competent)
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Gene transfer to animal cells
downstream of the polyhedrin promoter. The foreign
gene is cloned in another plasmid between two Tn7
repeats and introduced into the bacmid-containing
bacteria, which also contains a third plasmid expressing Tn7 transposase. Induction of transposase
synthesis results in the site-specific transposition of
the transgene into the bacmid, generating a recombinant baculovirus genome that can be isolated for
transfection into insect cells. Transposition of the
transgene into the bacmid interrupts the lacZ gene,
allowing recombinant bacterial colonies to be identified by blue–white screening.
Although baculoviruses productively infect insect
cells, they can also be taken up by mammalian cells,
although without producing progeny virions. The
number of mammalian cell lines reported to be transduced by baculoviruses is growing (reviewed by Kost
& Condreay 1999) and this suggests that recombinant baculoviruses could be developed as vectors for
gene therapy. A number of baculovirus-borne transgenes have been expressed using constitutive promoters, such as the cytomegalovirus immediate
early promoter (Hoffman et al. 1995) and the Rous
sarcoma virus promoter (Boyce & Bucher 1996).
More recently, cell lines have been generated that
are stably transformed with baculovirus vectors,
although it is not clear whether the viral genome
has integrated in these cells (Condreay et al. 1999).
Herpesvirus vectors
The herpesviruses are large dsDNA viruses that
include EBV (discussed above) and the HSVs (e.g.
HSV-I, varicella zoster). Most HSVs are transmitted
without symptoms (varicella zoster virus is exceptional) and cause prolonged infections. Unlike EBV,
which is used as a replicon vector, HSV-I has been
developed as a transduction vector. Viral replication
can occur in many cell types in a wide range of
species if the genome is introduced by transfection,
but HSV vectors are particularly suitable for gene
therapy in the nervous system, because the virus is
remarkably neurotropic. As with other large viruses,
recombinants can be generated in transfected cells
by homologous recombination, and such vectors
may be replication-competent or helper-dependent
(Marconi et al. 1996). Alternatively, plasmid-based
193
amplicon vectors can be constructed, which carry
only those cis-acting elements required for replication and packaging. These require packaging systems
to provide the missing functions in trans (e.g.
Stavropoulos & Strathdee 1998). Therapeutic use of
herpesvirus vectors has been limited, but a number
of genes have been successfully transferred to neurons in vivo (e.g. Boviatsis et al. 1994, Lawrence et al.
1995). Generally, transgene expression is transient,
although prolonged expression has been observed
in some neuronal populations (see reviews by Vos
et al. 1996, Simonato et al. 2000). Note that HSV
is also transmitted across neuronal synapses during lytic infections, a phenomenon that can be exploited to trace axon pathways (Norgren & Lehman
1998).
Retrovirus vectors
Retroviruses are RNA viruses that replicate via a
dsDNA intermediate. The infection cycle involves
the precise integration of this intermediate into the
genome of the host cell, where it is transcribed
to yield daughter genomes that are packaged into
virions.
Retroviruses have been developed as vectors for a
number of reasons (reviewed by Miller 1992). First,
certain retroviruses are acutely oncogenic because
they carry particular genes that promote host cell
division. Investigation of such viruses has shown
that these viral oncogenes are in fact gain-of-function
derivatives of host genes, proto-oncogenes, which are
normally involved in the regulation of cell growth.
In most cases, the viral oncogenes are found to be
expressed as fusions with essential viral genes, rendering the virus replication-defective. These acute
transforming retroviruses therefore demonstrate the
natural ability of retroviruses to act as replicationdefective gene-transfer vectors. Secondly, most retroviruses do not kill the host, but produce progeny
virons over an indefinite period. Retroviral vectors
can therefore be used to make stably transformed
cell lines. Thirdly, viral gene expression is driven by
strong promoters, which can be subverted to control
the expression of transgenes. In the case of murine
mammary-tumour virus, transcription from the viral
promoter is inducible by glucocorticoids, allowing
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CHAPTER 10
transgenes controlled by this promoter to be
switched on and off (Lee et al. 1981, Scheidereit et al.
1983). Fourthly, some retroviruses, such as amphotropic strains of murine leukaemia virus (MLV),
have a broad host range, allowing the transduction
of many cell types. Finally, retroviruses make efficient and convenient vectors for gene transfer because
the genome is small enough for DNA copies to be
manipulated in vitro in plasmid cloning vectors, the
vectors can be propagated to high titres (up to 108
pfu/ml) and the efficiency of infection in vitro can
approach 100%.
The major disadvantage of oncoretroviral vectors
is that they only productively infect dividing cells,
which limits their use for gene-therapy applications
(Miller et al. 1990, Roe et al. 1993). However, lentiviruses, such as human immunodeficiency virus
(HIV), are more complex retroviruses that have
the ability to infect non-dividing cells (Lewis &
Emmerman 1994). These were initially developed
as vectors for the stable transduction of cells displaying CD4 (Poznansky et al. 1991, Shimada et al.
1991, Buchschacher & Panganiban 1992, Parolin
et al. 1994), but recent advances in lentiviral vector
design provide improved safety and allow the transduction of multiple cell types (Naldini et al. 1996;
reviewed by Naldini 1998).
Before discussing the development of retroviral
vectors, it is necessary to briefly describe the genome
structure and the molecular biology of the infection
cycle (for a comprehensive account, see Weiss et al.
1985). A typical retroviral genome map is shown
in Fig. 10.8. The infection cycle begins when the
viral envelope interacts with the host cell’s plasma
membrane, delivering the particle into the cell. The
capsid contains two copies of the RNA genome, as
well as reverse transcriptase/integrase. Thus, immediately after infection, the RNA genome is reversetranscribed to produce a cDNA copy. This is a
complex process involving two template jumps,
with the result that the terminal regions of the RNA
genome are duplicated in the DNA as long terminal
repeats (LTRs). The DNA intermediate then integrates into the genome at an essentially random
site (there may be some preference for actively transcribed regions). The integrated provirus has three
genes (gag, pol and env). The gag gene encodes a viral
structural protein, pol encodes the reverse transcriptase and integrase and the env gene encodes viral
envelope proteins. Viral genomic RNA is synthesized
by transcription from a single promoter located in
the left LTR and ends at a polyadenylation site in the
right LTR. Thus, the full-length genomic RNA is
shorter than the integrated DNA copy and lacks the
duplicated LTR structure. The genomic RNA is
capped and polyadenylated, allowing the gag gene to
be translated (the pol gene is also translated by readthrough, producing a Gag–Pol fusion protein, which
is later processed into several distinct polypeptides).
Some of the full-length RNA also undergoes splicing,
eliminating the gag and pol genes and allowing the
downstream env gene to be translated. Two copies of
the full-length RNA genome are incorporated into
each capsid, which requires a specific cis-acting
packaging site termed ψ. The reverse transcriptase/
integrase is also packaged.
Strategies for vector construction
Most retroviral vectors are replication-defective,
because removal of the viral genes provides the
maximum capacity for foreign DNA (about 8 kb).
ψ
LTR
U3 R U5
LTR
gag
pol
env
PB(-)
ψ
R U5
U3 R U5
PB(+)
gag
pol
env
U3 R polyA
Fig. 10.8 Generic map of an oncoretrovirus genome. Upper figure shows the structure of an integrated provirus, with long
terminal repeats (LTRs) comprising three regions U3, R and U5, enclosing the three open reading frames gag, pol and env. Lower
figure shows the structure of a packaged RNA genome, which lacks the LTR structure and possesses a poly(A) tail. PB represents
primer binding sites in the viral replication cycle, and ψ is the packaging signal. The small circles represent splice sites.
POGC10 9/11/2001 11:04 AM Page 195
Gene transfer to animal cells
Only the cis-acting sites required for replication and
packaging are left behind. These include the LTRs
(necessary for transcription and polyadenylation of
the RNA genome as well as integration), the packaging site ψ, which is upstream of the gag gene, and
‘primer-binding sites’, which are used during the
complex replication process. The inclusion of a small
portion of the gag coding region improves packaging
efficiency by up to 10-fold (Bender et al. 1987).
Deleted vectors can be propagated only in the presence of a replication-competent helper virus or a
packaging cell line. The former strategy leads to the
contamination of the recombinant vector stock with
non-defective helper virus. Conversely, packaging
lines can be developed where an integrated provirus
provides the helper functions but lacks the cis-acting
sequences required for packaging (Mann et al. 1983).
Many different retroviruses have been used to
develop packaging lines and, since these determine
the type of envelope protein inserted into the virion
envelope, they govern the host range of the vector
(they are said to pseudotype the vector). Packaging
lines based on amphotropic MLVs allow retroviral
gene transfer to a wide range of species and cell
types, including human cells (e.g. see Cone &
Mulligan 1984, Danos & Mulligan 1988). It is still
possible for recombination to occur between the
vector and the integrated helper provirus, resulting
in the production of wild-type contaminants. The
most advanced ‘third-generation’ packaging lines
limit the extent of homologous sequence between
the helper virus and the vector and split up the
coding regions so that up to three independent
crossover events are required to form a replicationcompetent virus (e.g. see Markowitz et al. 1988).
The simplest strategy for the high-level constitutive expression of single genes in retroviral vectors is
to delete all coding sequences and place the foreign gene between the LTR promoter and the viral
polyadenylation site. Alternatively, an internal
heterologous promoter can be used to drive transgene expression. However, many investigators have
reported interference between the heterologous promoter and the LTR promoter (e.g. see Emmerman
& Temin 1984, Wu et al. 1996). Yu et al. (1986)
addressed this problem by devising self-inactivating
vectors, containing deletions in the 3′ LTR, which
are copied to the 5′ LTR during vector replication,
195
thus inactivating the LTR promoter while leaving
internal promoters intact. This strategy also helps to
alleviate additional problems associated with the
LTR promoter: (i) that adjacent endogenous genes
may be activated following integration; and (ii) that
the entire expression cassette may be inactivated by
DNA methylation after a variable period of expression in the target cell (Naviaux & Verma 1992).
Since retroviral vectors are used for the production of stably transformed cell lines, it is necessary
to co-introduce a selectable marker gene along
with the transgene of interest. The expression of two
genes can be achieved by arranging the transgene
and marker gene in tandem, each under the control
of a separate promoter, one of which may be the LTR
promoter. This leads to the production of full-length
and subgenomic RNAs from the integrated provirus.
Alternatively, if the first gene is flanked by splice
sites, only a single promoter is necessary, because
the RNA is spliced in a manner reminiscent of the
typical retroviral life cycle, allowing translation of
the downstream gene (Cepko et al. 1984). Vectors in
which the downstream gene is controlled by an
internal ribosome entry site have also been used
(e.g. Dirks et al. 1993, Sugimoto et al. 1994; see
Box 13.3). Since the viral replication cycle involves
transcription and splicing, an important consideration for vector design is that the foreign DNA must
not contain sequences that interfere with these
processes. For example, polyadenylation sites downstream of the transgene should be avoided, as these
will cause truncation of the RNA, blocking the replication cycle (Miller et al. 1983). Retroviruses also
remove any introns contained within the transgene
during replication (Shimotohno & Temin 1982).
Sindbis virus and Semliki forest virus
(alphaviruses)
The alphaviruses are a family of enveloped viruses
with a single-strand positive-sense RNA genome.
One of the advantages of using such RNA viruses
for gene transfer is that integration into the host
genome is guaranteed never to occur. Alphavirus
replication takes place in the cytoplasm, and produces a large number of daughter genomes, allowing very high-level expression of any transgene
(reviewed by Berglung et al. 1996). To date, these
POGC10 9/11/2001 05:57 PM Page 196
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CHAPTER 10
Poly A
Subgenomic
promoter to drive
expression of
inserted genes
vectors (Berglung et al. 1996, Lundstrom 1997). For
example, replication-competent vectors have been
constructed in which an additional subgenomic
promoter is placed either upstream or downstream
of the capsid polyprotein gene. If foreign DNA is
introduced downstream of this promoter, the replicase protein produces two distinct subgenomic
RNAs, one corresponding to the transgene. Such
insertion vectors tend to be unstable and have been
largely superseded by replacement vectors in which
the capsid polyprotein gene is replaced by the transgene. The first 120 bases of both the Sindbis and the
SFV structural polyprotein genes include a strong
enhancer of protein synthesis, which significantly
increases the yield of recombinant protein (Frolov
& Schlesinger 1994, Sjoberg et al. 1994). This is
downstream of the translational initiation site, so in
many vectors this enhancer region is included so
that the foreign gene is expressed as an N-terminal
fusion protein. This can result in extremely high
levels of recombinant protein synthesis – up to 50%
of the total cellular protein.
Both plasmid replicon and viral transduction
vectors have been developed from the alphavirus
genome. A versatile Sindbis replicon vector, pSinRep5,
is currently marketed by Invitrogen (Fig. 10.9). The
vector is a plasmid containing bacterial backbone
elements, the Sindbis replicase genes and packaging
Pac I
Not I
Xho I
Xba I
Mlu I
Pml I
Sph I
Stu I
Apa I
are the only animal viruses with a replication cycle
based solely on RNA to be extensively developed as
expression vectors. Sindbis virus (Xiong et al. 1989)
and Semliki Forest virus (SFV) (Liljestrom & Garoff
1993) have been the focus of much of this research.
These display a broad host range and cell tropism,
and mutants have been isolated with reduced cytopathic effects (Agapov et al. 1998, Frolov et al. 1999;
reviewed by Schlesinger & Dubensky 1999).
The wild-type alphavirus comprises two genes: a
5′ gene encoding viral replicase and a 3′ gene encoding a polyprotein from which the capsid structural
proteins are autocatalytically derived. Since the
genome is made of RNA, it can act immediately as a
substrate for protein synthesis. However, because
protein synthesis in eukaryotes is dependent on
mRNAs possessing a specialized 5′ cap structure,
only the replicase gene is initially translated. The
replicase protein produces a negative-sense complementary strand, which in turn acts as a template
for the production of full-length daughter genomes.
However, the negative strand also contains an internal promoter, which allows the synthesis of a subgenomic positive-sense RNA containing the capsid
polyprotein gene. This subgenomic RNA is subsequently capped and translated.
A number of different strategies have been used
to express recombinant proteins using alphavirus
PSG
3’ restriction sites for
linearization prior to
in vitro transcription
Am
pic
illin
IE1
Co
en
es
PS
S P6
pSinRep5
lG
ra
tu
uc
str
Non
Nonstructural
genes (nsP1–4)
for in vitro
replication of
recombinant
RNA
1–
4 (R
epli
case)
SP6 promoter
for synthesis of
in vitro RNA
transcripts
Packaging signal
allows recombinant
RNA to be packaged
into virus
Fig. 10.9 The Sindbis virus-based
vector pSinRep5. (Reproduced with
permission from Invitrogen.)
POGC10 9/11/2001 11:04 AM Page 197
Gene transfer to animal cells
site, and an expression cassette featuring a Sindbis
subgenomic promoter, a multiple cloning site and
a polyadenylation site. There is an SP6 promoter
upstream of the replicase genes and expression cassette for generating full-length in vitro transcripts.
There is a second set of restriction sites downstream
from the polylinker, allowing the vector to be linearized prior to in vitro transcription. Foreign DNA
is cloned in the expression cassette, the vector is
linearized and transcribed and the infectious recombinant Sindbis RNA thus produced is transfected into cells, from which recombinant protein
can be recovered. An alternative strategy is to place
the entire alphavirus genome under the control of a
standard eukaryotic promoter, such as SV40, and to
transfect cells with the DNA. In this case, the DNA
is expressed in the nucleus and the recombinant
vector RNA is exported into the cytoplasm. Such
DNA vectors have been described for Sindbis and
SFV (Herweijer et al. 1995, Johanning et al. 1995,
Berglund et al. 1996). Transduction is a more suitable
delivery procedure for gene therapy applications,
and in this case a replicon vector, such as pSinRep5,
is co-introduced with a defective helper plasmid
supplying the missing structural proteins. This facilitates one round of replication and packaging and
the production of recombinant viral particles, which
can be isolated from the extracellular fluid. In the
original packaging system, the replicon and helper
vectors could undergo recombination to produce
moderate amounts of contaminating wild-type
virus. This problem has been addressed by supplying
the structural-protein genes on multiple plasmids
(Smerdou & Liljestrom 1999) and by the development of complementary cell lines producing the
structural proteins (Polo et al. 1999).
Vaccinia and other poxvirus vectors
Vaccinia virus is closely related to variola virus, the
agent responsible for smallpox. A worldwide vaccination programme using vaccinia virus resulted in the
elimination of smallpox as an infectious disease. The
success of the programme raised hopes that recombinant vaccinia viruses, carrying genes from other
pathogens, could be used as live vaccines for other
infectious diseases. This expectation appears to have
been borne out by recent successes, as discussed below.
197
The poxviruses have a complex structure and a
large double-stranded linear DNA genome (up to
300 kb). Unusually for a DNA virus, the poxviruses
replicate in the cytoplasm of the infected cell rather
than its nucleus. Part of the reason for the large
genome and structural complexity is that the virus
must encode and package all its own DNA replication and transcription machinery, which most DNA
viruses ‘borrow’ from the host cell nucleus. The
unusual replication strategy and large size of the
vaccinia genome make the design and construction
of expression vectors more complex than for other
viruses. Since the virus normally packages its own
replication and transcription enzymes, recombinant
genomes introduced into cells by transfection are
non-infectious. Recombinant viruses are therefore
generated by homologous recombination, using a
targeting plasmid transfected into virus-infected cells.
More recently, direct ligation vectors have been
developed, and these are transfected into cells
containing a helper virus to supply replication and
transcription enzymes in trans (Merchlinsky et al.
1997). Recombinant vectors can be identified by
hybridization to the large viral plaques that form on
permissive cells. However, the efficiency of this process can be improved by various selection regimes.
In one strategy, the transgene is inserted into the
viral Tk gene and negative selection, using the thymidine analogue 5-bromodeoxyuridine, is carried out
to enrich for potential recombinants (Mackett et al.
1982). In another, the transgene is inserted into the
viral haemagglutinin locus. If chicken erythrocytes
are added to the plate of infected cells, wild-type
plaques turn red whereas the recombinant plaques
remain clear (Shida 1986). Since vaccinia vectors
have a high capacity for foreign DNA, selectable
markers, such as neo (Franke et al. 1985), or
screenable markers, such as lacZ (Chakrabarti et al.
1985) or gusA (Carroll & Moss 1995), can be cointroduced with the experimental transgene to
identify recombinants.
Transgene expression usually needs to be driven
by an endogenous vaccinia promoter, since transcription relies on proteins supplied by the virus.
The highest expression levels are provided by late
promoters such as P11, allowing the production of
up to 1 µg of protein per 106 cells, but other promoters
such as P7.5 and 4b are used, especially where
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CHAPTER 10
expression early in the infection cycle is desired
(Bertholet et al. 1985, Cochran et al. 1985). A synthetic late promoter, whose use allows up to 2 µg of
protein to be produced per 106 cells, has also been
developed (Lundstrom 1997). Since the cytoplasm
lacks not only host transcription factors but also the
nuclear splicing apparatus, vaccinia vectors cannot
be used to express genes with introns. Furthermore,
the sequence TTTTTNT must be removed from all
foreign DNA sequences expressed in vaccinia vectors, since the virus uses this motif as a transcriptional terminator. A useful binary expression system
has been developed, in which the transgene is driven
by the bacteriophage T7 promoter and the T7 polymerase itself is expressed in a vaccinia vector under
the control of a vaccinia promoter (Fuerst et al.
1986). Initially, the transgene was placed on a plasmid vector and transfected into cells infected with
the recombinant vaccinia virus, but higher expression levels (up to 10% total cellular protein) can be
achieved using two vaccinia vectors, one carrying
the T7-driven transgene and the other expressing
the polymerase (Fuerst et al. 1987).
In an early demonstration that vaccinia virus could
be used to express antigens from other infectious
agents, Smith, G.L. et al. (1983) replaced the vaccinia
Tk locus with a transgene encoding hepititis B surface
antigen (HBSAg). The transgene was cloned in a
plasmid containing the vaccinia Tk gene, interrupted
by one of the vaccinia early promoters. This plasmid
was transfected into vaccinia-infected monkey cells,
and recombinant vectors carrying the transgene were
selected using 5-bromodeoxyuridine. Cells infected
with this virus secreted large amounts of HBSAg into
the culture medium, and vaccinated rabbits rapidly
produced high-titre antibodies to HBSAg. Similarly,
vaccinia viruses expressing the influenza haemagglutinin gene were used to immunize hamsters and
induce resistance to influenza (Smith, G.E. et al.
1983). Recombinant vaccinia viruses have been
constructed expressing a range of important antigens, including HIV and HTLV-III envelope proteins
Table 10.4 Summary of major expression systems used in animal cells.
System
Host
Major applications
Non-replicating plasmid vectors
No selection
Dominant selectable markers
DHFR/methotrexate
Many cell lines
Many cell lines
CHO cells
Transient assays
Stable transformation, long-term expression
Stable transformation, high-level expression
Plasmids with viral replicons
SV40 replicons
BPV replicons
EBV replicons
COS cells
Various murine
Various human
High-level transient expression
Stable transformation (episomal)
Stable transformation (episomal), library
construction
293 cells
Various mammalian
Various mammalian
Insects
Various mammalian
Various mammalian and avian
ES cells
Non-dividing cells, mammalian
Various mammalian
Various mammalian
Transient expression
In vivo transfer
In vivo transfer
High-level transient expression
In vivo transfer
Stable transformation
Transgenic mice (Chapter 11)
In vivo transfer
High-level transient expression
High-level transient expression
Viral transduction vectors
Adenovirus E1 replacement
Adenovirus amplicons
Adeno-associated virus
Baculovirus
Oncoretrovirus
Lentivirus
Sindbis, Semliki Forest virus
Vaccinia virus
POGC10 9/11/2001 11:04 AM Page 199
Gene transfer to animal cells
(Chakrabarti et al. 1986, Hu et al. 1986). In many
cases, recombinant vectors have been shown to
provide immunity when administered to animals
(reviewed by Moss 1996). For example, monkeys
infected with recombinant vaccinia and canarypox
vectors have shown resistance to simian immunodeficiency virus (SIV) and HIV-2 (Hirsch et al.
1996, Myagkikh et al. 1996). Several recombinant
poxviruses have progressed to phase 1 clinical trials,
including a vaccine against the Epstein–Barr major
membrane antigen (Gu et al. 1996). The immunization of wild foxes, using ‘spiked’ meat, with a recombinant vaccinia virus expressing the rabies-virus
glycoprotein appears to have eliminated the disease
from Belgium (Brochier et al. 1991).
199
Summary of expression systems
for animal cells
A variety of gene transfer and expression systems
have been discussed in this chapter, and Table 10.4
provides a summary of these systems and their major
applications. Many factors influence the expression
of foreign genes in animal cells, and an understanding of these factors allows transgene expression to be
controlled. For the production of recombinant proteins, it is often appropriate to maximize transgene
expression, and principles for achieving this are
shown in Box 10.2. In other cases, it may be desirable to switch the transgene on and off, using
inducible expression systems. We explore these considerations in more detail in Chapter 13.
Box 10.2 Construct design for high-level transgene expression in animal cells
Many of the expression systems discussed in
this chapter are used in experiments where the
production of recombinant protein is the ultimate
aim. In such cases, it is appropriate for transgene
expression to be maximized. Although the different
expression systems vary in their total potential
yield, in terms of vector design and experimental
methodology, the following considerations should
apply when high-level expression is required.
The use of a strong and constitutive promoter
Very active promoters provide the highest levels of
transgene expression. In viral vectors, transgenes are
often expressed under the control of the strongest
endogenous promoters, e.g. the baculovirus
polyhedrin promoter, the adenoviral E1 promoter
and the vaccinia virus p7.5 promoter. Certain
viruses contain strong promoters and enhancers that
function in a wide range of cell types, and several of
these have been subverted for use in plasmid vectors.
The elements most commonly used in mammalian
cells are the SV40 early promoter and enhancer
(Mulligan & Berg 1981b), the Rous sarcoma virus
long-terminal-repeat promoter and enhancer
(Gorman et al. 1982a) and the human
cytomegalovirus immediate early promoter (Boshart
et al. 1985). Although these function widely, they are
not necessarily active in all mammalian cells, e.g.
the SV40 promoter functions poorly in the human
embryonic kidney line 293 (Gorman et al. 1989).
The inclusion of an intron
The presence of an intron in a eukaryotic expression
unit usually enhances expression. Evidence for the
positive effect of introns accumulated in the early
years of cDNA expression in animal cells, although
there are also many studies in which efficient gene
expression was obtained in the absence of an intron
(see Kaufman 1990). Nevertheless, most mammalian
expression vectors in current use incorporate a
heterologous intron, such as the SV40 small t-antigen
intron or the human growth-hormone intron, or
modified hybrid introns that match the consensus
splice donor and acceptor-site sequences. Note that
introns may not be used in some expression systems,
such as vaccinia virus. The presence of an intron is
very important in constructs that are to be expressed
in transgenic animals (see Box 11.1 for more
discussion).
continued
POGC10 9/11/2001 11:04 AM Page 200
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CHAPTER 10
Box 10.2 continued
The inclusion of a polyadenylation signal
Polyadenylation signals (terminators) are required in
eukaryotic genes to generate a defined 3′ end to the
mRNA. In most cases, this defined end is extended by
the addition of several hundred adenosine residues
to generate a poly(A) (polyadenylate) tail. This tail is
required for the export of mRNA into the cytoplasm,
and also increases its stability. In the absence of such
a site, the level of recombinant protein produced in
transformed cells can fall by as much as 90%
(Kaufman 1990). Poly(A) sites from the SV40 early
transcription unit or mouse b-globin gene are often
incorporated into mammalian expression vectors.
The removal of unnecessary
untranslated sequence
ATG
The sequence around the translational initiation
site should conform to Kozak’s consensus, which is
defined as 5′-CCRCCAUGG-3′ (Kozak 1986, 1999).
Of greatest importance is the purine at the −3
position (identified as R) and the guanidine at
position +4. The adenosine of the AUG initiation
codon (underlined) is defined as position +1, and
the immediately preceding base is defined as
Igκ Leader
P CMV
BGH
myc epitope
6xHis
TAA
pA
f1
or
i
40
Psv
pSecTag2
Zeoc
in
T7
Optimization of the transgene for
translational efficiency
Sfi I*
Asc I
Hind III
Asp718
Kpn I
BamH I
BstX I
EcoR I*
Pst I*
EcoR V
BstX I
Not I
Xho I
Dra II
Apa I
Eukaryotic mRNAs comprise a coding region (which
actually encodes the gene product) bracketed by
untranslated regions (UTRs) of variable lengths.
Both the 5′ and 3′ UTRs can influence gene
expression in a number of ways (Kozak 1999). For
example, the 5′ UTR may contain one or more AUG
codons upstream of the authentic translational start
site, and these are often detrimental to translational
initiation. The 3′ UTR may contain regulatory
elements that control mRNA stability (e.g. AU-rich
sequences that reduce stability have been identified
(see Shaw & Kamen 1986)). Furthermore, both the
5′ and 3′ UTRs may be rich in secondary structure,
which prevents efficient translation. In animal
systems, UTR sequences are generally removed from
transgene constructs to maximize expression.
li
cil
pi
Am
n
pA
40
SV
ColE1
continued
POGC10 9/11/2001 11:04 AM Page 201
Gene transfer to animal cells
201
Box 10.2 continued
position −1. The expression of foreign genes in
animals can also be inefficient in some cases due
to suboptimal codon choice, which reflects the fact
that different organisms prefer to use different
codons to specify the same amino acid. If a transgene
contains a codon that is commonly used in the source
organism but rarely used in the host, translation
may pause at that codon due to the scarcity of the
corresponding transfer RNA (tRNA). This will reduce
the rate of protein synthesis and may even lead to
truncation of the protein or frame-shifting. It may
therefore be beneficial to ‘codon-optimize’
transgenes for the expression host.
The incorporation of a targeting signal
If the goal of an expression study is to recover large
amounts of a functional eukaryotic protein, it is
necessary to consider whether that protein needs to
be post-translationally modified in order to function
correctly. For example, many proteins intended for
therapeutic use require authentic glycosylation
patterns not only for correct function, but also to
prevent an immune response in the patient. Since
specific types of modification occur in particular cell
compartments, it is necessary to consider strategies
for targeting the recombinant protein to the correct
compartment to ensure that it is appropriately
modified. Proteins that need to be glycosylated, for
example, must be targeted to the secretory pathway
using a signal peptide. Many mammalian expression
vectors are available for this purpose, and they
incorporate heterologous signal peptides. The figure
opposite shows the Invitrogen vector pSecTag2,
which incorporates a sequence encoding the murine
immunoglobulin light-chain signal peptide for highefficiency targeting to the secretory pathway. Note
also that the C terminus of the recombinant protein is
expressed as a fusion to two different epitope tags to
facilitate protein purification (see p. 76).
POGC11 9/11/2001 11:15 AM Page 202
CH A P T E R 11
Genetic manipulation of animals
Introduction
The genetic manipulation of animals has revolutionized our understanding of biology, by making it
possible to test gene expression and function at the
whole-animal level. Gene-transfer techniques can
be used to produce transgenic animals, in which every
cell carries new genetic information, as well as
designer mutants with specific preselected modifications to the genome. The whole animal is the
ultimate assay system in which to investigate gene
function, particularly for complex biological processes, such as development.
In the case of plants, gene transfer to tissues or
cultured cells is often the first step in the production
of a transgenic organism, since whole fertile plants
can regenerate from such cells and explants under
the appropriate culture conditions. Two fundamental
differences between plants and animals make this
strategy impossible in animals. First, animal cells
become progressively restricted in terms of developmental potency as development proceeds, which
means that differentiated animal cells are normally
unable to fully dedifferentiate and recapitulate the
developmental programme.* Secondly, in most
animals, the somatic cells and germ cells (the cells
that give rise to gametes) separate at an early developmental stage. Therefore, the only way to achieve
germ-line transformation in animals is to introduce
* Recently, adult haemopoietic stem cells have been shown to
contribute to fetal blood-cell development when injected into
the blastocysts of mouse embryos (Geiger et al. 1998). Under
similar circumstances, bone marrow and neural stem cells
have been shown to contribute to specific developing tissues
(Petersen et al. 1999, Clarke et al. 2000). This suggests that
mammalian development may be more plastic than previously assumed, and could open the way for novel routes to
animal transgenesis.
DNA into totipotent cells prior to the developmental
stage at which the germ line forms.
In most cases, this involves introducing DNA
directly into the developing oocyte, egg or early
embryo. In mice, it is also possible to use cultured
embryonic stem cells (ES cells), which are derived
from the preimplantation embryo and can contribute to all the tissues of the developing animal
(including the germ line) if introduced into a host
embryo at the correct developmental stage. These
cells are also remarkably amenable to homologous
recombination, which allows them to be used for
gene targeting, the accurate replacement of a segment of the endogenous genome with a homologous
segment of exogenous DNA. Gene targeting can be
used to replace endogenous genes with a completely
non-functional copy (a null allele) or to make subtle
changes, both allowing the function of the endogenous gene to be tested. The same technology can be
used for the opposite purpose, i.e. to replace a
mutant allele with a functional copy.
While rapid progress has been made with a range
of model organisms, especially the fruit fly Drosophila
melanogaster, the mouse and, more recently, the
African clawed frog, Xenopus, great potential exists
in the production of transgenic farm animals with
improved or novel traits. The technology for introducing DNA into animals such as chickens, pigs,
cattle and sheep is still in its infancy and is much
less efficient than that of mice. Furthermore, it has
proved impossible to isolate amenable ES cells from
any species except mice and chickens. The desire to
generate transgenic livestock has driven research
in a different direction, that of nuclear transfer.
Although differentiated animal cells are developmentally restricted, their nuclei still contain all the
genetic information required to recapitulate the
whole of development. Dolly, the first sheep produced following nuclear transfer from a differentiated somatic cell to an enucleated egg, has opened
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Genetic manipulation of animals
the way for animal cloning and the rapid production
of élite transgenic herds.
Genetic manipulation of mammals
Methods for producing transgenic mice
The ability to introduce DNA into the germ line of
mice is one of the greatest achievements of the
twentieth century and has paved the way for
the transformation of other mammals. Genetically
modified mammals have been used not only to study
gene function and regulation, but also as bioreactors
producing valuable recombinant proteins, e.g. in
their milk. Several methods for germ-line transformation have been developed, all of which require the
removal of fertilized eggs or early embryos from
donor mothers, brief culture in vitro and then their
return to foster-mothers, where development continues to term. These methods are discussed below
and summarized in Fig. 11.1.
203
Pronuclear microinjection
Direct microinjection of DNA was the first strategy
used to generate transgenic mice. Simian virus 40
(SV40) DNA was injected into the blastocoele
cavities of preimplantation embryos by Jaenisch and
Mintz (1974). The embryos were then implanted
into the uteri of foster-mothers and allowed to
develop. The DNA was taken up by some of the
embryonic cells and occasionally contributed to the
germ line, resulting in transgenic mice containing
integrated SV40 DNA in the following generation.
Transgenic mice have also been recovered following
the injection of viral DNA into the cytoplasm of the
fertilized egg (Harbers et al. 1981).
The technique that has become established is the
injection of DNA into one of the pronuclei of the egg
(reviewed by Palmiter & Brinster 1986). The technique is shown in Fig. 11.2. Just after fertilizaton,
the small egg nucleus (female pronucleus) and the
large sperm nucleus (male pronucleus) are discrete.
Attach DNA
to sperm
DNA transfection
Adult cells
Unfertilized
mouse egg
Nuclear
replacement
Embryonic stem
(ES) cells
Recombinant
retroviral infection
Fertilized egg
DNA
microinjection
Cell transfer
Early cleavage
Blastocyst
Fig. 11.1 Summary of methods for producing
transgenic mammals.
Embryonic
development
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CHAPTER 11
Recombinant retroviruses
Fig. 11.2 Pronuclear microinjection of a fertilized mouse
egg. The two pronuclei are visible, and the egg is held using
a suction pipette. The DNA is introduced through a fine
glass needle. (Photograph courtesy of Roberta Wallace,
Roslin Institute.)
Since the male pronucleus is larger, this is usually
chosen as the target for injection. About 2 pl of DNA
solution is transferred into the nucleus through a
fine needle, while the egg is held in position with a
suction pipette. The injected embryos are cultured in
vitro to the morula stage and then transferred to
pseudopregnant foster-mothers (Gordon & Ruddle,
1981). The procedure requires specialized microinjection equipment and considerable dexterity from
the handler. The exogenous DNA may integrate
immediately or, less commonly, may remain extrachromosomal for one or more cell divisions. Thus
the resulting animal may be transgenic or may be
chimeric for transgene insertion. The technique
is reliable, although the efficiency varies, so that
5–40% of mice developing from manipulated eggs
contain the transgene (Lacy et al. 1983). However,
once the transgene is transmitted through the germ
line, it tends to be stably inherited over many generations. The exogenous DNA tends to form head-to-tail
arrays prior to integration, and the copy number
varies from a few copies to hundreds. The site of
integration appears random and may depend on the
occurrence of natural chromosome breaks. Extensive
deletions and rearrangements of the genomic DNA
often accompany transgene integration (Bishop &
Smith 1989).
As discussed in Chapter 10, recombinant retroviruses provide a natural mechanism for stably
introducing DNA into the genome of animal cells.
Retroviruses are able to infect early embryos and ES
cells (see below), so recombinant retroviral vectors
can be used for germ-line transformation (Robertson
et al. 1986). An advantage over the microinjection
technique is that only a single copy of the retroviral
provirus is integrated, and the genomic DNA surrounding the transgenic locus generally remains
intact. The infection of preimplantation embryos
with a recombinant retrovirus is technically straightforward and, once the infected embryos are implanted
in the uterus of a foster-mother, can lead to germline transmission of the transgene. However, there
are also considerable disadvantages to this method,
including the limited amount of foreign DNA that
can be carried by the virus, the possible interference
of viral regulatory elements with the expression of
surrounding genes and the susceptibility of the virus
to de novo methylation, resulting in transgene silencing (see Box 13.2). The founder embryos are always
chimeric with respect to transgene integration
(reviewed by Jaenisch 1988). Retroviral transduction
is therefore not favoured as a method for generating
fully transgenic animals, but it is useful for generating transgenic sectors of embryos. For example, the
analysis of chicken-limb buds infected with recombinant retroviruses has allowed many of the genes
involved in limb development to be functionally
characterized (see review by Tickle & Eichele 1994).
Transfection of ES cells
ES cells are derived from the inner cell mass of the
mouse blastocyst and thus have the potential to
contribute to all tissues of the developing embryo
(Evans & Kaufman 1981, Martin 1981). The ability
of ES cells to contribute to the germ line was first
demonstrated by Bradley et al. (1984) and requires
culture conditions that maintain the cells in an undifferentiated state (Joyner 1998). Since these cells can
be serially cultured, like any other established cell
line, DNA can be introduced by transfection or
viral transduction and the transformed cells can
be selected using standard markers, as discussed in
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Genetic manipulation of animals
Chapter 10. This is an important advantage, since
there is no convenient way to select for eggs or
embryos that have taken up foreign DNA, so,
instead, each potential transgenic mouse must be
tested by Southern-blot hybridization or the polymerase chain reaction (PCR) to confirm transgene
integration. ES cells are also particularly efficient at
carrying out homologous recombination (see below),
so, depending on the design of the vector, DNA introduced into ES cells may integrate randomly or may
target and replace a specific locus.
Whichever strategy is chosen, the recombinant
ES cells are then introduced into the blastocoele of a
host embryo at the blastocyst stage, where they mix
with the inner cell mass. This creates a true chimeric
embryo (i.e. an embryo comprising cells from different sources). The contribution of ES cells to the germ
line can thus be confirmed using visible markers.
Most ES cell lines in common use are derived from
mouse strain 129, which has the dominant coat
colour agouti. A popular strategy is to use host
embryos from a mouse strain such as C57BL/6J,
which has a recessive black coat colour. Colonization of the embryo by vigorous ES cells can be
substantial, generating chimeras with patchwork
coats of black and agouti cell clones. If the ES cells
have contributed to the germ line, mating chimeric
males with black females will generate heterozygous
transgenic offspring with the agouti coat colour,
confirming germ-line transmission of the foreign
DNA. Most ES cells in use today are derived from
male embryos, resulting in a large sex bias towards
male chimeras (McMahon & Bradley, 1990). This
is desirable because male chimeras sire many more
offspring than females.
Gene targeting with ES cells
Pronuclear microinjection and retroviral transfer
are useful for the addition of DNA to the mouse
genome. However, in many cases it is more desirable
to replace endogenous gene sequences with exogenous DNA, since this would allow the introduction
of specific mutations into any preselected gene. In
yeast, gene targeting by homologous recombination
occurs with high efficiency (Chapter 9). In contrast,
when the first gene-targeting experiments in animal
cells were carried out in the 1980s, only a very low
205
frequency of targeted recombination was achieved.
These experiments involved the correction of mutations in selectable markers such as neo, which had
been introduced into cell lines as transgenes by
standard methods (e.g. Thomas et al. 1986). Smithies
et al. (1985) were the first to demonstrate targeting
of an endogenous gene. They introduced a modified
β-globin gene containing the bacterial marker supF
into a human fibroblast × mouse erythroleukaemia
cell line and screened large numbers of potential
recombinants by reisolating the modified locus, using
supF as a cloning tag. This experiment demonstrated
that the frequency of homologous recombination
was up to 1000-fold lower than that of random integration. Targeting occurs with significantly higher
efficiency in certain cell lines, including mouse ES
cells. The combination of pluripotency, amenability
for in vitro manipulation and capacity for homologous recombination makes ES cells uniquely
suitable for the generation of targeted mutant mice,
i.e. mice carrying the same mutation in every cell
and transmitting it through the germ line. Gene
targeting in ES cells was first achieved by Thomas
and Capecchi (1987), who disrupted the hprt gene
with the neo marker and selected recombinant cells
using either G418 or 6-thioguanine, a toxic guanine
analogue that is only incorporated into DNA if
the nucleotide salvage pathway is functional (see
p. 178). Doetschman et al. (1987) also successfully
targeted the hprt locus, although they used a mutant
recipient cell line and repaired the locus with homologous DNA, subsequently selecting on HAT medium
(see p. 177).
Design of targeting vectors
Targeting vectors are specialized plasmid vectors,
which promote homologous recombination when
introduced into ES cells. This is achieved by the
inclusion of a homology region, i.e. a region that is
homologous to the target gene, allowing the targeting vector to synapse with the endogenous DNA.
Both the size of the homology region and the level
of sequence identity have been shown to play an
important role in the efficiency of gene targeting
(Hasty et al. 1991b, Deng & Capecchi 1992, Te Riele
et al. 1992). Recombination is also more efficient if
the vector is linearized prior to transfection.
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Most gene targeting experiments have been used
to disrupt endogenous loci, resulting in targeted null
alleles (this strategy is often termed ‘gene knockout’).
Two types of targeting vector have been developed
for this purpose: insertion vectors and replacement
(or transplacement) vectors (Thomas & Capecchi
1987; Fig. 11.3). Insertion vectors are linearized
within the homology region, resulting in the insertion of the entire vector into the target locus. This
type of vector disrupts the target gene but leads
to a duplication of the sequences adjacent to the
selectable marker. This is not always a desirable
configuration, since duplication of the target sequences
can lead to a subsequent homologous recombination event that restores the wild-type genotype
(Fiering et al. 1995). Replacement vectors are
designed so that the homology region is collinear
with the target. The vector is linearized outside the
homology region prior to transfection, resulting in
crossover events in which the endogenous DNA is
replaced by the incoming DNA. With this type of
vector, only sequences within the homology region
(not the vector backbone) are inserted, so, to achieve
gene knockout, the homology region itself must
be interrupted. Insertion and replacement vectors
are thought to be equally efficient, but replacement vectors have been used in the majority of
knockout experiments. In both cases, however, it
is possible for transcription to occur through the
targeted locus, producing low amounts of RNA.
This may be spliced in such a way as to remove the
targeted exon, resulting in a residual amount of
functional protein (e.g. Dorin et al. 1994, Dahme
et al. 1997).
while in replacement vectors the marker must
interrupt the homology region. The neo marker has
been most commonly used, allowing transformed
ES cells to be selected using G418. However, other
dominant markers are equally applicable (e.g. see
Von Melchner et al. 1992), and Hprt can be used in
combination with hprt− mutant ES cells (Matzuk
et al. 1992).
The use of a single marker fails to discriminate
between targeted cells and those where the construct has integrated randomly. This problem can be
addressed by combined positive–negative selection
using neo and the herpes simplex virus (HSV) Tk
gene (Mansour et al. 1988). If the neo marker is used
to interrupt the homology region in a replacement
vector, transformed cells can be selected using G418.
The HSV Tk gene is placed outside the homology
region, such that it is inserted by random integration
but not by homologous recombination. Therefore,
cells that have undergone homologous recombination will survive in the presence of the toxic thymidine analogues ganciclovir or FIAU,* while in those
cells containing randomly integrated copies of the
Tk gene, the analogues will be incorporated into the
DNA, resulting in cell death.
A different strategy is to make expression of the
neo gene dependent on homologous recombination
(e.g. see Schwartzberg et al. 1989, Mansour et al.
1993). Another alternative to positive–negative
selection, which is used in many laboratories, is
simply to screen large numbers of G418-resistant
transfected cells by PCR to identify genuine recombinants (Hogan & Lyons 1988, Zimmer & Gruss
1989). Screening can be carried out relatively quickly
without recourse to cloning.
Selection strategy
The first gene-targeting experiments involved the
selectable Hprt locus, which is present on the X
chromosome, allowing targeted events to be selected
in male ES cells without the requirement for homozygosity. The first non-selectable genes to be targeted
were int-2 (also known as fgf-3) (Mansour et al.
1988) and the oncogene c-abl (Schwartzberg et al.
1989). In each case, it was necessary to include a
selectable marker in the targeting vector to identify
transformed cells. In the case of insertion vectors
this is placed anywhere on the vector backbone,
Introducing subtle mutations
While gene targeting has often been used to disrupt
and hence inactivate specific endogenous genes by
introducing large insertions, more refined approaches
can be used to generate subtle mutations. The
precise effects of minor deletions or point mutations
cannot be assessed using the simple targeting strategies discussed above, which necessarily leave the
* 1-(2-deoxy-2-fluoro-β-d-arabinofuranosyl)-5-iodouracil.
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Genetic manipulation of animals
207
Targeting vector
2
1
×
2
×
3
Homologous target
4
Integration (ends-in)
1
2
Targeting vector
1
3*
3*
2
3
4
1
4
×
2
3
4
Transplacement (ends-out)
1
4
Fig. 11.3 Structure and integration mechanisms of the two major types of targeting vector, (a) insertion and (b) transplacement.
Regions of homology between the vector and target are designated by the same number.
Second replacement vector
(containing subtle mutation)
Intrachromosomal
recombination
1
2
1
3*
×
×
2
3
2
3*
1
4
×
4
4
Second round
transplacement
1
2
3*
4
Mutation
1
2
3*
4
Fig. 11.4 Procedure for the introduction of subtle mutations using (a) an insertion vector and (b) two transplacement vectors.
Regions of homology between the vector and target are designated by the same number. An asterisk indicates the position of a
point mutation in the vector, which is to be introduced at the target locus.
selectable marker and, in some cases, the entire
targeting vector integrated at the target site. Furthermore, many investigators have reported that
the strong promoters used to drive marker-gene
expression can affect the regulation of neighbouring
genes, in some cases up to 100 kb away from the targeted locus (Pham et al. 1996). Two major strategies
for the introduction of subtle mutations have been
devised, each involving two rounds of homologous
recombination. These are the ‘hit-and-run’ strategy,
involving a single insertion vector, and the ‘tagand-exchange’ strategy (Fig. 11.4), involving two
replacement vectors. Strategies involving Cre recombinase have also been developed and are discussed
on p. 257.
The ‘hit-and-run’ or ‘in–out’ strategy (Hasty et al.
1991a, Valancius & Smithies 1991) involves the
use of an insertion vector carrying two selectable
markers, such as neo and Tk. The insertion event is
positively selected using G418. As discussed above,
the use of an insertion vector results in duplication of
the homology region in the targeted clone, although
in this case the homology region derived from the
vector is modified to contain the desired subtle
mutation. The success of this strategy relies on a
second intrachromosomal homologous recombination event, which replaces the endogenous allele
with the mutant and deletes the markers, allowing the second-round recombinants to survive in
the presence of ganciclovir. However, the second
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CHAPTER 11
homologous recombination event occurs at a very
low frequency and, in 50% of cases, restores the
locus to its original wild-type configuration.
The ‘tag-and-exchange’ strategy also requires two
homologous recombination events, but in this case
two replacement-type vectors are used. For example, Moore et al. (1995) demonstrated the principle
using neo for positive selection and HSV Tk for negative selection. The first ‘tag’ vector was designed to
mutate the target gene by inserting a large cassette
containing the selectable markers. This event was
positively selected with G418. The second ‘exchange’
vector introduced the desired mutation and eliminated
the selectable markers, allowing the second-round
recombinants to be selected for the absence of Tk.
Applications of genetically modified mice
Applications of transgenic mice
Transgenic mice, i.e. mice containing additional
transgenes, as opposed to those with targeted mutations, have been used to address many aspects of
gene function and regulation. A vast literature has
accumulated on this subject, and genes concerning
every conceivable biological process have been
investigated (e.g. see Houdebine 1997). As well as
their use for basic scientific investigation, transgenic
mice can be used for more applied purposes, such
as models for human disease and the production
of valuable pharmaceuticals. Many mouse models
for human diseases have been generated by gene
knockout (see below), but gain-of-function models
have also been generated by adding transgenes. For
example, much information concerning the pathology of prion diseases has arisen from the study of
transgenic mice expressing mutant prion transgenes (reviewed by Gabizon & Taraboulos 1997).
Transgenic mice expressing oncogenes have been
extensively used to study cancer (reviewed by
Macleod & Jacks 1999).
For illustrative purposes, we now consider some
early experiments that demonstrated how transgenic mice can be used for the analysis of gene
function and regulation, but also highlighted some
limitations of the transgenic approach. Brinster et al.
(1981) constructed plasmids in which the promoter
of the mouse metallothionein-1 (MMT) gene was
fused to the coding region of the HSV Tk gene.
The thymidine kinase (TK) enzyme can be assayed
readily and provides a convenient reporter of MMT
promoter function. The endogenous MMT promoter
is inducible by glucocorticoid hormones and heavy
metals, such as cadmium and zinc, so it was envisaged
that the hybrid transgene, MK (metallothioneinthymidine kinase), would be similarly regulated.
The gene was injected into the male pronucleus of
fertilized eggs, which were then incubated in vitro in
the presence or absence of cadmium ions (Brinster
et al. 1982). As expected, TK activity was found to be
induced by the metal. By making a range of deletions
of mouse sequences upstream of the MMT promoter
sequences, the minimum region necessary for inducibility was localized to a stretch of DNA 40–180
nucleotides upstream of the transcription-initiation
site. Additional sequences that potentiate both basal
and induced activities extended to at least 600 bp
upstream of the transcription-initiation site. The
mouse egg was therefore being used in the same way
as transfected cell lines, to dissect the activity of a functional promoter (Gorman et al. 1982b; see Box 10.l).
The same MK fusion gene was injected into
embryos, which were raised to transgenic adults
(Brinster et al. 1981). Most of these mice expressed
the MK gene and in such mice there were from one
to 150 copies of the gene. The reporter activity was
inducible by cadmium ions and showed a tissue
distribution very similar to that of metallothionein
itself (Palmiter et al. 1982b). Therefore these experiments showed that DNA sequences necessary for
heavy-metal induction and tissue-specific expression
could be functionally dissected in both eggs and
transgenic mice. For unknown reasons, there was
no response to glucocorticoids in either the egg or
the transgenic-mouse experiments.
In a dramatic series of experiments, Palmiter et al.
(1982a) fused the MMT promoter to the rat growthhormone gene. This hybrid gene (MGH) was constructed using the same principles as the MK fusion.
Of 21 mice that developed from microinjected eggs,
seven carried the MGH fusion gene and six of these
grew significantly larger than their littermates. The
mice were fed zinc to induce transcription of the MGH
gene, but this did not appear to be absolutely necessary, since they showed an accelerated growth rate
before being placed on the zinc diet. Mice containing
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209
Examples of increased, decreased or even totally
extinguished expression were found. In some, but
not all, cases, the changes in expression correlated
with changes in methylation of the gene sequences
(Palmiter et al. 1982b). These results provided the
first examples of two complex phenomena, position
effects (Box 11.1) and de novo transgene silencing
(Box 13.2), which often affect integrated transgenes.
Yeast artificial chromosome (YAC)
transgenic mice
Fig. 11.5 Transgenic mouse containing the mouse
metallothionein promoter fused to the rat growth-hormone
gene. The photograph shows two male mice at about
10 weeks old. The mouse on the left contains the MGH gene
and weighs 44 g; his sibling without the gene weighs 29 g.
In general, mice that express the gene grow two to three times
as fast as controls and reach a size up to twice the normal.
(Photograph by courtesy of Dr R.L. Brinster.)
high copy numbers of the MGH gene (20–40 copies
per cell) had very high concentrations of growth
hormone in their serum, some 100–800 times above
normal. Such mice grew to almost double the weight
of littermates at 74 days old (Fig. 11.5).
The similarities between the tissue distribution of
normal MMT expression and that of the hybrid
transgenes encouraged the hope that transgenic
mice would provide a general assay for functionally
dissecting DNA sequences responsible for tissuespecific or developmental regulation of a variety of
genes. However, there were also some unexpected
findings. For example, independently derived transgenic mice carrying the MK transgene showed
significant variations in the levels and patterns of
transgene expression. Furthermore, while transgenic founders transmitted the construct to their
progeny as expected, when reporter activity was
assayed in these offspring the amount of expression
could be very different from that in the parent.
Studies of the MMT promoter and others have
demonstrated the principle that transgenes with
minimal flanking sequences tend not to be expressed
in the same manner as the corresponding endogenous gene. In many cases, it has also been shown
that authentic patterns and levels of protein expression occur only when the intact gene is used, and
this can span tens or hundreds of kilobase pairs
of DNA (Box 11.1). The transfer of large DNA
segments to the mouse genome has been achieved
by transformation with yeast artificial chromosome
(YAC) vectors. Jakobovits et al. (1993) were the first
to report transformation of ES cells with a YAC
vector, via fusion with yeast sphaeroplasts. The vector contained the entire human HPRT locus, nearly
700 kb in length. The disadvantage of this method is
that the endogenous yeast chromosomes were cointroduced with the vector. Alternative strategies
involve isolation of the vector DNA by pulsed-field
gel electrophoresis (p. 10), followed by introduction
of the purified YAC DNA into mouse eggs by pronuclear microinjection or transfection into ES cells.
The latter technique is more suitable because microinjection involves shear forces that break the DNA
into fragments. YAC transfer to ES cells has been
achieved by lipofection, as discussed in Chapter 10.
YAC transgenics have been used to study gene
regulation, particularly by long-range regulatory
elements, such as locus-control regions (reviewed
by Lamb & Gerhart 1995). They have also been used
to introduce the entire human immunoglobulin locus
into mice, for the production of fully humanized
antibodies (Mendez et al. 1997). It is also possible to
introduce chromosomes and chromosome fragments
into ES cells, using a technique called microcellmediated fusion. This involves the prolonged mitotic
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Box 11.1 Position effects
Independently derived transgenic animals and plants
carrying the same expression construct often show
variable levels and patterns of transgene expression.
In many cases, such variation is dependent on the
site of transgene integration, and this phenomenon
has been termed the position effect (reviewed by
Wilson et al. 1990). Position effects result from
the influence of local regulatory elements on the
transgene, as well as the architecture of the
surrounding chromatin. For example, an integrated
transgene may come under the influence of a local
enhancer, resulting in the alteration of its expression
profile to match that of the corresponding
endogenous gene. The position dependence of
the phenomenon has been demonstrated in mice
by isolating the entire transgenic locus from such
an anomalous line and microinjecting it into the
pronuclei of wild-type eggs, resulting in ‘secondary’
transgenic lines with normal transgene expression
profiles (Al-Shawi et al. 1990). Position effects are
also revealed by enhancer-trap constructs, which
contain a minimal promoter linked to a reporter
gene (O’Kane & Gehring 1987; see Chapter 13).
Unlike the specific influences of nearby regulatory
elements, chromatin-mediated position effects are
generally non-specific and repressive. They reflect
the integration of the transgene into a chromosomal
region containing repressed chromatin
(heterochromatin). The molecular features of
heterochromatin, including its characteristic
nucleosome structure, deacetylated histones and,
in many cases, hypermethylated DNA, spread into
the transgene, causing it to be inactivated (Huber
et al. 1996, Pikaart et al. 1998). In some cases,
variegated transgene expression has been reported
due to cell-autonomous variations in the extent of
this spreading process (reviewed by Heinkoff 1990).
Negative chromosomal position effects can be
troublesome in terms of achieving desirable
transgene expression levels and patterns; thus
a number of different strategies have been used
to combat them.
Incorporating dominantly acting
transcriptional control elements
Certain regulatory elements are thought to act as
master-switches, regulating the expression of genes
or gene clusters by helping to establish an open
chromatin domain. The locus control region (LCR)
of the human b-globin gene cluster is one example
(Forrester et al. 1987). Transgenic mice carrying
a human b-globin transgene driven by its own
promoter show a low frequency of expression and, in
those mice that do express the transgene, only a low
level of the mRNA is produced (e.g. Magram et al.
1985, Townes et al. 1985). However, inclusion of the
LCR in the expression construct confers high-level
and position-independent expression (Grosveld et al.
1987). There is evidence that LCRs induce chromatin
remodelling over large distances. For example, the
murine immunoglobulin heavy-chain LCR has been
shown to induce histone deacetylation in a linked
c-myc gene (Madisen et al. 1998). This suggests
that LCRs could protect against position effects by
converting heterochromatin to open euchromatin at
the site of transgene integration (Festenstein et al.
1996, Milot et al. 1996). The interested reader can
consult several comprehensive reviews of LCR
research (Bonifer 1999, Grosveld 1999, Li et al.
1999).
Using boundary elements/matrix
attachment regions
Boundary elements (insulators) are sequences that
can block the activity of enhancers when placed
between the enhancer and a test transgene driven by
a minimal promoter. For example, an ‘A element’ with
insulator activity is found upstream of the chicken
lysozyme gene. This inhibits the activity of a reporter
gene when interposed between the promoter and an
upstream enhancer, but not when placed elsewhere
in the construct (Stief et al. 1989). However,
by flanking the entire construct with a pair of
continued
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211
Box 11.1 continued
A elements, the transgene is protected from
chromosomal position effects (Stief et al. 1989). This
protective effect works not only in cell lines, but also
in transgenic animals (McKnight et al. 1992) and
plants (Mlynarova et al. 1994). Many boundary
elements are associated with matrix-attachment
regions (MARs), sequences dispersed throughout the
genome that attach to the nuclear matrix, dividing
chromosomes into topologically independent loops
(reviewed by Spiker & Thompson 1996). It is
therefore possible that transgenes flanked by such
elements are maintained in an isolated chromatin
domain into which heterochromatin cannot spread.
However, not all boundary elements are associated
with MARs (e.g. see Mirkovitch et al. 1984). Similarly,
some MARs do not function as boundary elements
but as facilitators of gene expression (e.g. Van der
Geest & Hall 1997).
Using large genomic transgenes
Conventional transgenes generally comprise
complementary DNAs (cDNAs) or intronless
‘minigenes’ expressed under the control of viral
promoters or cell-type-specific regulatory elements.
Such transgenes are highly sensitive to position
effects. Over the last few years, there has been an
increasing appreciation that the regulation of
eukaryotic gene expression is far more complex
and involves much more upstream and downstream
DNA than previously thought (reviewed by Bonifer
1999, 2000). The correct, high-level expression
of transgenes is favoured by the use of genomic
constructs that include introns and large amounts
arrest of cultured human cells, using an inhibitor
such as colchicine. Eventually, the nucleus breaks
up into vesicles containing individual chromosomes,
which can be rescued as microcells comprising a
nuclear vesicle surrounded by a small amount of
cytoplasm and a plasma membrane (Fournier &
Ruddle 1977). Transgenic mice have been generated
using ES cells that were fused to human microcells,
and evidence for germ-line transmission and ex-
of flanking sequence from the source gene (e.g. see
Bonifer et al. 1990, Lien et al. 1997, Nielsen et al.
1998). Such constructs are likely to include multiple
enhancers, dominant regulatory elements, such as
LCRs, and boundary elements, which all act together
to protect the transgene from position effects.
Dominantly acting transgenes
(transgene rescue)
Some conventional transgenes, including b-globin
and a-fetoprotein (Chada et al. 1986, Kollias et al.
1986, Hammer et al. 1987) are very sensitive to
position effects and de novo silencing. Other genes
appear to be less sensitive to these phenomena, e.g.
immunoglobulin and elastase (Storb et al. 1984,
Swift et al. 1984, Davis & MacDonald 1988).
Although the reason for this is not clear, the less
sensitive transgenes are assumed to in some way
induce or define an open chromatin domain. In some
cases, such sequences have been used to protect
more susceptible transgenes from negative position
effects by introducing the two transgenes
simultaneously (e.g. see Clark et al. 1992).
Site-specific integration
Site-specific recombination systems (see Chapter 13)
can be used to introduce transgenes into a locus
known to lack negative position effects, if a target
site for the recombinase can be introduced at such
a locus. The Cre-loxP system has been used to this
effect in mammalian cells (see Fukushige & Sauer
1992).
pression of the human chromosome was obtained
(Tomizuka et al. 1997).
Applications of gene targeting
Since the first reports of gene targeting in ES cells, an
ever-increasing number of targeted mutant mice
have been produced. These have been discussed
in several comprehensive reviews (Brandon et al.
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1995a,b,c, Soriano 1995, Muller 1999) and a number of Internet databases have been established to
keep track of the results (see Sikorski & Peters 1997).
The phenotypes of homozygous, null mutant mice
provide important clues to the normal function of
the gene. Some gene knockouts have resulted in
surprisingly little phenotypic effect, much less severe
than might have been expected. For example, myoD,
whose expression in transfected fibroblasts causes
them to differentiate into muscle cells, and which
was therefore a good candidate as a key regulator of
myogenesis, is not necessary for development of a
viable animal (Rudnicki et al. 1992). Similarly, the
retinoic acid γ receptor is not necessary for viable
mouse development in knockout mice (Lohnes et al.
1993), even though this receptor is a necessary
component of the pathway for signalling by retinoids
and has a pattern of expression quite distinct from
other retinoic acid receptors in embryos. Such observations have prompted speculation that genetic
redundancy may be common in development, and
may include compensatory up-regulation of some
members of a gene family when one member is
inactivated. An example of this may be the upregulation of myf-5 in mice lacking myoD (Rudnicki
et al. 1992). Gene knockouts have also been used as
mouse models of human diseases, such as cystic
fibrosis, β-thalassaemia and fragile X syndrome
(reviewed by Bedell et al. 1997; see Chapter 14).
While most gene-targeting experiments in mice
have been used to introduce mutations into genes
(either disruptive insertional mutations or subtle
changes), the scope of the technique is much wider.
The early gene-targeting experiments demonstrated
that this approach could also be used to correct
mutated genes, with obvious applications in gene
therapy. Homologous recombination has also been
used to exchange the coding region of one gene for
that of another, a strategy described as ‘gene knockin’. This has been used, for example, to test the
ability of the transcription factors Engrailed-1 and
Engrailed-2 to compensate for each other’s functions.
Hanks et al. (1995) replaced the coding region of the
engrailed-1 gene with that of engrailed-2, and showed
that the engrailed-1 mutant phenotype could be
rescued. A more applied use of gene knock-in is
the replacement of parts of the murine immunoglobulin genes with their human counterparts,
resulting in the production of humanized antibodies
in transgenic mice (Moore et al. 1995). The Cre-loxP
site-specific recombinase system has been used extensively in ES cells to generate mice in which conditional or inducible gene targeting is possible and to
produce defined chromosome deletions and translocations as models for human disease. We shall
discuss the many applications of Cre-loxP and other
site-specific recombinase systems in Chapter 13.
Other mammals and birds
Traditional techniques
The three major routes for producing transgenic mice
have also been used in other mammals and birds,
particularly in farm animals. The efficiency of each
procedure is much lower than in mice. Pronuclear
microinjection in mammals such as sheep and cows,
for example, typically results in less than 1% of the
injected eggs giving rise to transgenic animals. Added
to this, the recovery of eggs from donor animals and
the reimplantation of transformed eggs into fostermothers is a less efficient procedure and requires, at
great expense, a large number of donors and recipients. The eggs themselves are also more difficult to
manipulate – they are very delicate and tend to be
opaque. It is often necessary to centrifuge the eggs in
order to see the pronuclei. In chickens, it is possible
to remove eggs just after fertilization and microinject
DNA into the cytoplasm of the germinal disc, where
the male and female pronuclei are to be found.
However, it is not possible to return the manipulated
eggs to a surrogate mother, so they must be cultured
in vitro. Using this procedure, Love et al. (1994)
obtained seven chicks, equivalent to about 5% of
the eggs injected, that survived to sexual maturity.
One cockerel transmitted the transgene to a small
proportion of his offspring, indicating that he was
chimeric for transgene integration.
The use of retroviruses to produce transgenic
chickens has been reported by Bosselman et al.
(1989). These investigators injected a replicationdefective recombinant reticuloendotheliosis virus
carrying the neo gene into laid eggs and found
that approximately 8% of male birds carried vector
sequences. The transgene was transmitted through
the germ line in a proportion of these birds and
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was stably expressed in 20 transgenic lines. It is
necessary for the nuclear envelope to break down
for most retroviral infections (lentiviruses such as
human immunodeficiency virus (HIV) are exceptional) and, since this only occurs during mitosis,
most retroviruses are unable to infect non-dividing
cells (Chapter 10). The nuclear envelope also breaks
down during meiosis, and this was exploited by
Chan et al. (1998) to produce transgenic cattle
following the injection of replication-defective retroviral vectors into the perivitelline space of isolated
bovine oocytes. Retroviral integration occurred during the second meiotic division, resulting in the
production of a number of transgenic offspring.
Remarkably, the same technique has recently been
used to generate the first ever transgenic primate, a
rhesus monkey named ANDi (Chan et al. 2001).
In primates, the technique was inefficient. Two
hundred and twenty-four oocytes were injected to
produce one live transgenic monkey; a number of
further transgenic fetuses failed to develop to term.
After more than a decade of research, it has also
proved impossible to derive reliable ES cell lines from
any domestic species other than mice and, more
recently, chickens (Pain et al. 1999, Prelle et al.
1999)*. However, there have been great advances
in the isolation and transfection of primordial germ
cells (PGCs), the embryonic cells that give rise to
gametes. These can be transfected directly, or cultured as embryonic germ cells (EG cells), which are
morphologically very similar to ES cells and could
provide a route for the direct transformation of the
germ line (Resnick et al. 1992). Chicken PGCs have
been isolated from the germinal crescent, infected
with a recombinant retrovirus and replaced in the
embryo, leading to the development of chimeric
birds producing transgenic offspring (Vick et al.
1993). Mammalian PGCs have also been transformed, although it has been difficult to persuade the
cells to contribute to the germ line once introduced
into the host animal (Labosky et al. 1994).
Intracytoplasmic sperm injection
The injection of sperm heads directly into the cytoplasm of the egg (intracytoplasmic sperm injection
* Human ES cells and Human ES cell lines are available.
213
(ICSI)) can overcome infertility in humans. It has
been shown that sperm heads bind spontaneously to
naked plasmid DNA in vitro, suggesting that sperm
injections could be used to achieve transformation.
This was demonstrated by Perry et al. (1999), who
mixed mouse sperm with plasmid DNA carrying
the gene for green fluorescent protein (GFP). These
sperm were injected into unfertilized oocytes, and a
remarkable 94% of the resulting embryos showed
GFP activity. Random transfer of these embryos to
pseudopregnant females resulted in development
to term, and in about 20% of cases the mice were
transgenic. This method could be adaptable to other
animals. Rhesus monkey oocytes fertilized in the
same manner gave rise to a number of embryos with
GFP activity, but this only lasted until the blastula
stage, suggesting that there was no stable integration. However, several monkeys developed to term,
showing that the procedure was compatible with
normal development (Chan et al. 2000).
Nuclear transfer technology
The failure of traditional transgenesis techniques to
yield routine procedures for the genetic modification
of mammals other than mice has driven researchers
in search of other methods. Fifty years ago, Briggs
and King (1952) established the principle of nuclear
transfer in amphibians by transplanting nuclei from
the blastula of the frog Rana pipens to an enucleated
egg, obtaining a number of normal embryos in the
process. In Xenopus laevis, nuclei from various types
of cell in the swimming tadpole can be transplanted
to an egg that has been UV-irradiated to destroy the
peripheral chromosomes, and similar results are
obtained (reviewed by Gurdon 1986, 1991). The
important principle here is that, while animal cells
become irreversibly committed to their fate as
development proceeds, the nuclei of most cells still
retain all the genetic information required for the
entire developmental programme and can, under
appropriate circumstances, be reprogrammed by the
cytoplasm of the egg to recapitulate development. In
all species, it appears that the earlier the developmental stage at which nuclei are isolated, the greater
their potential to be reprogrammed. Nuclear transplantation can be used to generate clones of animals
with the same genotype by transplanting many
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CHAPTER 11
Fig. 11.6 Megan and Morag, the first sheep produced by
nuclear transfer from cultured cells. Reproduced by kind
permission of the Roslin Institute, Edinburgh.
somatic nuclei from the same individual into a series
of enucleated eggs (King & Briggs 1956). This allows
animals with specific and desirable traits to be
propagated. If possible in mammals, this would have
obvious applications in farming.
Nuclear transfer in mammals has been practised
with success for the last decade, although rabbits
and farm animals, such as sheep, pigs and cows, are
far more amenable to the process than mice. In each
case, donor nuclei were obtained from the morula or
blastocyst-stage embryo and transferred to an egg or
oocyte from which the nucleus had been removed
with a pipette (Smith & Wilmut 1989, Willadsen
1989, Collas & Robl 1990, McLaughlin et al. 1990).
The donor nucleus can be introduced by promoting
fusion between the egg and a somatic cell. A brief
electric pulse is often used to achieve this, as it also
activates embryonic development by stimulating
the mobilization of calcium ions.
A major advance was made in 1995, when two
live lambs, Megan and Morag (Fig. 11.6), were produced by nuclear transfer from cultured embryonic
cells (Campbell et al. 1996). This demonstrated the
principle that mammalian nuclear transfer was possible using a cultured cell line. The same group later
reported the birth of Dolly (Fig. 11.7), following
nuclear transfer from an adult mammary epithelial
cell line (Wilmut et al. 1997). This was the first
mammal to be produced by nuclear transfer from a
differentiated adult cell, and aroused much debate
Fig. 11.7 Dolly and her lamb Bonnie. Dolly was the first
mammal to be generated by nuclear transfer from an
adult cell. Reproduced by kind permission of the Roslin
Institute, Edinburgh.
among both scientists and the public concerning the
possibility of human cloning (see Johnson 1998). It
was suggested that a critical factor in the success of
the experiment was the quiescent state of the cells
in culture, allowing synchronization between the
donor and recipient cell cycles. For the production of
Dolly, this was achieved by lowering the level of
serum in the culture medium, causing the cells to
withdraw from the cell cycle due to lack of growth
factors. However, the success rate was very low:
only one of 250 transfer experiments produced a
viable lamb. Similar transfer experiments have since
been carried out in mice, cows, pigs and goats
(Cibelli et al. 1998, Wakayama et al. 1998, Baguisi
et al. 1999, Polejaeva et al. 2000).
The success of nuclear transfer in domestic mammals provides a new route for the production of
transgenic animals. This involves the introduction
of DNA into cultured cells, which are then used as a
source of donor nuclei for nuclear transfer. Such a
cell-based strategy has many advantages over traditional techniques, such as microinjection, including
the ability to screen transformed cells for high-level
transgene expression prior to the nuclear-transfer
step. The production of a transgenic mammal by
nuclear transfer from a transfected cell line was first
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Genetic manipulation of animals
achieved by Schnieke et al. (1997), who introduced
the gene for human factor IX into fetal sheep fibroblasts and transferred the nuclei to enucleated eggs.
The resulting sheep, Polly, produces the recombinant
protein in her milk and can therefore be used as a
bioreactor (Chapter 14). More recently, McCreath
et al. (2000) succeeded in producing a transgenic
sheep by nuclear transfer from a somatic cell whose
genome had been specifically modified by gene
targeting. A foreign gene was introduced into the
COL1A1 locus and was expressed at high levels in
the lamb.
DNA transfer to other vertebrates
Gene transfer to Xenopus
Xenopus oocytes as a heterologous
expression system
Since Gurdon et al. (1971) first showed that oocytes
synthesized large amounts of globin after they had
been microinjected with rabbit globin mRNA, the
Xenopus oocyte expression system has been a valuable tool for expressing a very wide range of proteins
from plants and animals (Colman 1984). X. laevis is
an African clawed frog. Oocytes can be obtained
in large numbers by removal of the ovary of an
adult female. Each fully grown oocyte is a large cell
(0.8–1.2 mm diameter) arrested at first meiotic
prophase. This large cell has a correspondingly large
nucleus (called the germinal vesicle), which is located
in the darkly pigmented hemisphere of the oocyte.
Due to the large size of the oocytes, mRNA – either
natural or synthesized by transcription in vitro,
using phage-T7 RNA polymerase (Melton 1987) –
can be readily introduced into the cytoplasm or
nucleus by microinjection. This is achieved using a
finely drawn glass capillary as the injection needle,
held in a simple micromanipulator. DNA can also be
injected. The oocyte nucleus contains a store of
the three eukaryotic RNA polymerases, enough to
furnish the needs of the developing embryo at least
until the 60 000-cell stage. The RNA polymerases
are available for the transcription of injected exogenous DNA. Using this system, it has therefore been
possible to express complementary DNAs (cDNAs)
linked to a heat-shock promoter or to mammalian
215
virus promoters (Ballivet et al. 1988, Ymer et al.
1989, Swick et al. 1992). In addition, vaccinia virus
vectors (Chapter 10) can be used for gene expression
in the cytoplasm (Yang et al. 1991).
An important aspect of the oocyte expression
system is that recombinant proteins are usually
correctly post-translationally modified and directed
to the correct cellular compartment. For example,
oocytes translate a wide variety of mRNAs encoding secretory proteins, modify them and correctly
secrete them (Lane et al. 1980, Colman et al. 1981).
Foreign plasma-membrane proteins are generally
targeted to the plasma membrane of the oocyte,
where they can be shown to be functional. The first
plasma-membrane protein to be expressed in this
system was the acetylcholine receptor from the electric organ of the ray, Torpedo marmorata (Sumikawa
et al. 1981). Injected oocytes translated mRNA
extracted from the electric organ and assembled
functional multi-subunit receptor molecules in the
plasma membrane (Barnard et al. 1982). Following
this work, the oocyte has become a standard heterologous expression system for plasma-membrane
proteins, including ion channels, carriers and receptors. The variety of successfully expressed plasmamembrane proteins is very impressive. However,
there are examples of foreign channels and receptors
being non-functional in oocytes, either due to lack
of coupling to second-messenger systems in the
oocyte, incorrect post-translational modification, or
other reasons (reviewed in Goldin 1991).
Xenopus oocytes for functional expression cloning
Functional expression cloning using oocytes was
first developed by Noma et al. (1986), using a strategy
outlined in Fig. 11.8. The following example, the
cloning of the substance-K receptor, is illustrative. It
has been found that oocytes can be made responsive
to the mammalian tachykinin neuropeptide, substance K, by injecting an mRNA preparation from
bovine stomach into the oocyte cytoplasm. The preparation contains mRNA encoding the substance-K
receptor protein, which is evidently expressed as a
functional protein and inserted into the oocyte
membrane. Masu et al. (1987) exploited this property to isolate a cDNA clone encoding the receptor.
The principle was to make a cDNA library from
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CHAPTER 11
Tissue
Size
fractionation
Extraction
Affinity chromatography
Poly(A)+RNA
Microinjection
Poly(A)+RNA
Microinjection
Xenopus
oocyte
Assay for
functional
expression
Synthesis
cDNA library
In vitro transcription
and capping
Fig. 11.8 Strategy for functional expression cloning, using
Xenopus oocytes as a heterologous expression system.
stomach mRNA, using a vector in which the cDNA
was flanked by a promoter for the SP6 or T7 RNA
polymerase. This allowed in vitro synthesis of mRNA
from the mixture of cloned cDNAs in the library.
The receptor clone was identified by testing for
receptor expression following injection of synthetic
mRNA into the oocyte cytoplasm. Repeated subdivision of the mixture of cDNAs in the library led
to the isolation of a single cloned cDNA. The strategy described above can only be applied to cloning
single-subunit proteins, not proteins composed of
different subunits or proteins whose function in
oocytes requires more than one foreign polypeptide.
This limitation was overcome by Lubbert et al.
(1987), who used a hybrid depletion procedure to
clone a serotonin-receptor cDNA.
A prerequisite for using the oocyte in functional
expression cloning is a knowledge of the oocyte’s
own ion channels, carriers and receptors. Endogenous activity may mask or interfere with the soughtafter function (for a review, see Goldin 1991).
Transient gene expression in Xenopus embryos
Messenger RNA, synthesized and capped in vitro,
can be microinjected into dejellied Xenopus embryos
at the one- or two-cell stage. The mRNA is distributed among the descendants of the injected cells
and is expressed during early development. This
approach has been exploited very widely for examining the developmental effects resulting from the
overexpression of normal or altered gene products
(reviewed by Vize & Melton 1991).
DNA can be introduced into Xenopus embryos
in the same manner. However, unlike the situation
in mammals, where the injected DNA integrates
rapidly into the genome, exogenous DNA in Xenopus
persists episomally and undergoes extensive replication (Endean & Smithies 1989). Bendig and Williams
(1983) provide a typical example of this process.
They injected a recombinant plasmid carrying Xenopus
globin genes into the egg and showed that the
amount of plasmid DNA increased 50- to 100-fold
by the gastrula stage. In later development, the
amount of DNA per embryo decreased, and most
of the persisting DNA co-migrated with highmolecular-weight chromosomal DNA. This difference
between mammals and amphibians probably reflects their distinct modes of early development. In
mammals, cleavage divisions are slow and asynchronous. Gene expression occurs throughout early
development and supplies the embryo with the
proteins it requires at a steady rate. Conversely,
there is no transcription in the early Xenopus embryo
and yet the cleavage divisions are rapid and synchronous. DNA replication relies on stored maternal
gene products, so there is a stockpile of chromatin
assembly proteins and replication enzymes. Exogenous DNA injected into Xenopus eggs is therefore
assembled immediately into chromatin and undergoes replication in tune with the rapid DNA synthesis
already occurring in the nucleus (Leno & Laskey
1991). Etkin et al. (1987) have analysed the replication of a variety of DNAs injected into Xenopus
embryos. It was found that various plasmids
increase to different extents. This was not simply
related to the size of the plasmid, but also reflected
the presence of specific sequences that inhibited
replication. Replication has also been found to
depend upon the conformation and number of
molecules injected (Marini et al. 1989).
Transgenic Xenopus
DNA injected into early Xenopus embryos is expressed in a mosaic fashion during development,
regardless of the promoter used, which limits the use
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Genetic manipulation of animals
of this system for the analysis of gene expression and
function. Some of the DNA does become incorporated into the genome and may be transmitted
through the germ line (Rusconi and Schaffner
1981). However, integration occurs at a very low
frequency and, given the long generation interval
of Xenopus laevis (12–18 months from egg to
adult), this is not an efficient way to generate transgenic frogs.
A simple and efficient process for large-scale
transgenesis in Xenopus has become available only
in the last few years (Kroll & Amaya 1996). In this
technique, known as restriction-enzyme-mediated
integration (REMI), linearized plasmids containing
the transgene of interest are mixed with decondensed sperm nuclei and treated with limiting
amounts of a restriction enzyme to introduce nicks
in the DNA. The nuclei are then transplanted into
unfertilized Xenopus eggs, where the DNA is repaired,
resulting in the integration of plasmid DNA into the
genome. This technique allows the production of up
to 700 transgenic embryos per person per day, most
of which survive at least to the swimming-tadpole
stage. The decondensed nuclei are extremely fragile,
so careful handling and transplantation within
about 30 min are required for a good yield of normal
transgenic embryos. In some cases, viable transgenic adults have been derived from the tadpoles
and transgenic X. laevis lines have been established
(Bronchain et al. 1999, Marsh-Armstrong et al.
1999). A disadvantage of X. laevis is that the species
is tetraploid. Offield et al. (2000) have therefore
established transgenic lines of the closely related but
diploid species Xenopus tropicalis, which also has a
shorter generation interval than its tetraploid cousin.
Since Xenopus is used worldwide as a developmental model organism, transgenic Xenopus technology has been rapidly adopted in many laboratories
and is being used to examine (or in many cases
re-examine) the roles of developmental genes. Thus
far, the sophisticated tools used in transgenic mice
have not been applied to Xenopus, but this is only a
matter of time. Recently, an inducible expression
system based on the use of a Xenopus heat-shock
promoter was described, allowing inducible control
of the GFP gene. This system has been used to investigate Wnt signalling in early Xenopus development
(Wheeler et al. 2000). As discussed above, one of the
217
early successes in transgenic mouse methodology
was the expression of rat growth hormone, resulting
in transgenic mice up to twice the size of their nontransgenic siblings. The role of growth hormone in
amphibian metamorphosis has now been examined
by expressing Xenopus growth hormone in transgenic frogs. The transgenic tadpoles developed at the
same rate as control tadpoles, but typically grew to
twice the normal size (Huang & Brown 2000). After
metamorphosis, the transgenic frogs also grew much
more quickly than controls and showed skeletal
defects.
Gene transfer to fish
Fish transgenesis can be used to study gene function
and regulation, e.g. in model species, such as the
zebrafish (Danio rerio) and medaka (Oryzias latipes),
and to improve the traits of commercially important
species, such as salmon and trout. Gene-transfer
technology in fish has lagged behind that of mammals, predominantly due to the lack of suitable
regulatory elements to control transgene expression.
The first transgenic fish carried transgenes driven
by mammalian or viral regulatory elements, and
their performance varied considerably. For example,
attempts to express growth-hormone genes in trout
initially met with little success, and this may have
been due to the inability of fish cells to correctly process mammalian introns (Betancourt et al. 1993).
However, fish are advantageous assay systems for
several reasons, including their fecundity, the fact
that fertilization and development are external and
the ease with which haploid and uniparental diploid
embryos can be produced (Ihssen et al. 1990).
Like frogs, the injection of DNA into fish eggs and
early embryos leads to extensive replication and
expression from unintegrated transgenes, so that
fish, like frogs, can be used for transient expression
assays (Vielkind 1992). Some of the DNA integrates
into the genome, leading to germ-line transmission
and the production of transgenic fish lines (reviewed
by Iyengar et al. 1996). There has been recent
progress in the development of transgenic fish
with enhanced growth characteristics, particularly
through the use of expression constructs that are
derived from the same species (e.g. Rahman et al.
1998; reviewed by Dunham 1999). It is likely that
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transgenic fish will be the first genetically modified
animals to enter the food-chain.
DNA transfer to invertebrates
Transgenic flies
Drosophila P elements
P elements are transposable DNA elements that, under
certain circumstances, can be highly mobile in the
germ line of D. melanogaster. The subjugation of these
sequences as specialized vector molecules in Drosophila was a landmark in Drosophila genetics. Through
the use of P-element vectors, any DNA sequence can
be introduced into the genome of the fly.
P elements cause a syndrome of related genetic
phenomena called P–M hybrid dysgenesis (Bingham
et al. 1982, Rubin et al. 1982). Dysgenesis occurs
when males of a P (paternally contributing) strain
are mated with females of an M (maternally contributing) strain, but not when the reciprocal cross is
made. The syndrome predominantly affects the
germ line and induces a high rate of mutation and
frequent chromosomal aberrations, resulting in
abnormal (dysgenic) hybrid offspring. In extreme
cases, there is failure to produce any gametes at all.
Hybrid dysgenesis occurs because P strains
contain transposable genetic elements, P elements,
which are mobilized in the eggs of M-strain females
(eggs that are permissive for P-element transposition are described as ‘M-cytotype’). The P elements
do not cause dysgenesis in crosses within P strains,
because they are not mobilized in P-cytotype eggs.
This is because the P element encodes a repressor of
its own transposase, which prevents transposition.
When a sperm from a P-cytotype male fertilizes the
egg of an M-cytotype female, the absence of repressor in the egg results in temporary derepression of
the transposase, such that P-element transposition
occurs at a high frequency. The high rate of mutation characteristic of the dysgenesis syndrome
reflects the insertion of P elements into multiple
genetic loci.
Several members of the P-element family have
been cloned and characterized (O’Hare & Rubin
1983). The prototype is a 2.9-kb element, while
other members of the family appear to have arisen
by internal deletion events. The elements are characterized by perfect 31-bp inverted terminal repeats,
which are recognized by the transposase. The prototype element contains a single gene, comprising four
exons, encoding the transposase (a truncated version of the transposase may act as the repressor).
The transposase primary transcript is differentially
spliced in germ cells and somatic cells, such that
functional transposase is produced only in germ
cells. Laski et al. (1986) showed this clearly by
making a P-element construct in which the differentially spliced third intron was precisely removed.
This element showed a high level of somatic transposition activity. Naturally occurring short P elements
are generally defective, because they do not encode
functional transposase. However, they do possess
the inverted terminal repeats and can be activated
in trans by transposase supplied by a non-defective
P element in the same nucleus.
Spradling and Rubin (1982) devised an approach
for introducing P-element DNA into Drosophila
chromosomes. Essentially, a recombinant plasmid
comprising a 2.9 kb P element together with some
flanking Drosophila DNA sequences, cloned in the
pBR322 vector, was microinjected into the posterior
pole of M-cytotype embryos. The embryos were
injected at the syncytial blastoderm stage, when the
cytoplasm has not yet become partitioned into
individual cells (Fig. 11.9). The posterior pole was
chosen because this is where the germ line originates, and P-element DNA in this region was
expected to be incorporated into the genome in a
proportion of the germ cells.
A screen of progeny lines showed that P elements
had indeed integrated at a variety of sites in each of
the five major chromosomal arms, as revealed by
in situ hybridization to polytene chromosomes. Pelement integration occurred by transposition, not
by random integration. This was proved by probing
Southern blots of restricted DNA and showing that
the integrated P element was not accompanied by
the flanking Drosophila or pBR322 DNA sequences
present in the recombinant plasmid (Spradling
& Rubin 1982). The injected plasmid DNA must
therefore have been expressed at some level before
integration, so as to provide transposase.
These experiments showed that P elements could
transpose with a high efficiency from injected plasmids
POGC11 9/11/2001 11:15 AM Page 219
Genetic manipulation of animals
Plasma membrane
Female pronucleus
Male pronucleus
Chorion
(protective
coat)
Micropyle
(through which
sperm enters)
Fertilized egg
Germ plasm
Posterior
end
DNA injected
Anterior
end
9 synchronous nuclear divisions
Diploid cleavage nuclei
Syncytial
blastoderm
About 12 nuclei migrate to
the posterior pole where they
form germ line precursor cells
or pole cells
Pole cells
(germ line
precursors)
Blastoderm cell
Individual
blastoderm
cells formed
Fig. 11.9 Early embryogenesis of Drosophila. DNA injected
at the posterior end of the embryo just prior to pole-cell
formation is incorporated into germ-line cells.
into diverse sites in the chromosomes of germ cells.
At least one of the integrated P elements in each
progeny line remained functional, as evidenced by
the hypermutability it caused in subsequent crosses
to M-cytotype eggs.
Development of P-element vectors for gene transfer
Rubin and Spradling (1982) exploited their finding
that P elements can be artificially introduced in the
Drosophila genome. A possible strategy for using the
219
P element as a vector would be to attempt to identify
a suitable site in the 2.9 kb P-element sequence
where insertion of foreign DNA could be made without disrupting genes essential for transposition.
However, an alternative strategy was favoured. A
recombinant plasmid was isolated which comprised
a short (1.2 kb), internally deleted member of the Pelement family together with flanking Drosophila
sequences, cloned in pBR322. This naturally defective P element did not encode transposase (O’Hare
& Rubin 1983). Target DNA was ligated into the
defective P element. The aim was to integrate this
recombinant P element into the germ line of injected
embryos by providing transposase function in trans.
Two approaches for doing this were initially tested.
In one approach a plasmid carrying the recombinant P element was injected into embryos derived
from a P–M dysgenic cross, in which transposase
activity was therefore expected to be high. A
disadvantage of this approach was that frequent
mutations and chromosomal aberrations would
also be expected. In the other approach, the plasmid
carrying the recombinant P element was co-injected
with a plasmid carrying the non-defective 2.9 kb
element.
In the first experiments of this kind, embryos homozygous for the rosy mutation were microinjected
with a P-element vector containing a wild-type rosy
gene. Both methods for providing complementing
transposase were effective. Rosy+ progeny, recognized
by their wild-type eye colour, were obtained from
20–50% of injected embryos. The chromosomes of
these flies contained one or two copies of the integrated rosy transgene. The rosy gene is a particularly
useful genetic marker. It produces a clearly visible
phenotype: Rosy− flies have brown eyes instead of
the characteristic red colour of Rosy+ flies. The rosy
gene encodes the enzyme xanthine dehydrogenase,
which is involved in the production of a precursor of
eye pigments. The rosy gene is not cell-autonomous:
expression of rosy anywhere in the fly, for example in
a genetically mosaic fly developing from an injected
larva, results in a wild-type eye colour. Selectable
markers have been used instead of visible markers
to identify transformed flies. These include the
alcohol dehydrogenase gene adh (Goldberg et al.
1983) and neo (Steller & Pirrotta 1985). Other eyecolour markers have been used, including white and
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CHAPTER 11
Structure of P-element derivative: Carnegie 20
Insert DNA as required
Polylinker
IR
31 bp inverted repeat (IR)
7.2 kb rosy fragment
pUC8 vector
Lacks own transposase gene
Structure of helper P-element: pπ 25.7 wings clipped
23 bp of IR deleted
therefore cannot be transposed
0
1
2
3
IR
Encodes transposase
vermilion (Ashburner 1989, Fridell & Searles 1991),
as well as alternative visible markers, such as rough
(which restores normal eye morphology) and yellow
(which restores normal body pigmentation and is
particularly useful for scoring larvae) (Locket et al.
1992; Patton et al. 1992).
A simple P-element vector is shown in Fig. 11.10
(Rubin & Spradling 1983). It consists of a P element
cloned in the bacterial vector pUC8. Most of the P
element has been replaced by the rosy gene, but the
terminal repeats essential for transposition have
been retained. The vector includes a polylinker site
for inserting foreign sequences. Transposition of the
recombinant vector into the genome of injected
larvae is brought about by co-injecting a helper P
element, which provides transposase in trans but
which cannot transpose itself because of a deletion
in one of its terminal inverted repeats. Such an
element is referred to as a wings-clipped element
Fig. 11.10 P-element derivatives as a
vector system. (See text for details.)
(Karess & Rubin 1984). An alternative strategy is to
inject purified transposase protein (Kaufman & Rio
1991). The capacity of P-element vectors is large,
although increasing the size of the recombinant
element appears to reduce the transposition frequency. Inserts of over 40 kb have been successfully
introduced into flies (Haenlin et al. 1985) and this
has allowed the construction of cosmid libraries
using P-element vectors (Speek et al. 1988).
As well as their use for germ-line transformation,
P elements have been exploited for insertional mutagenesis, as cloning tags and as entrapment vectors
to detect genes and regulatory elements. Similar
applications have been applied to other transposable
elements, such as the Ac–Ds transposons of maize,
and to other gene-transfer systems, such as retroviruses and the T-DNA of Agrobacterium tumefaciens.
These diverse uses of gene-transfer vectors are discussed in Chapter 13.
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CHAPTER 12
Gene transfer to plants
Introduction
The expression of foreign genes introduced into
plants was first achieved in the early 1980s. In the
20 years following these initial successes, we have
witnessed a revolution in plant genetic engineering,
with the transformation of well over 100 different
plant species now a routine procedure. Our ability
to manipulate the plant genome has come about
through intensive research into vector systems based
on the soil bacterium Agrobacterium tumefaciens and
alternative strategies involving direct DNA transfer.
In addition, plant viruses have been developed as
versatile episomal vectors for high-level transient gene
expression. This research has immense biotechnological implications in the creation of plants with
useful genetically engineered characteristics, such as
insect resistance, herbicide tolerance and improved
nutritional value (Chapter 14).
A fundamental difference between animals and
plants is that organized, differentiated plant tissue
shows a high degree of developmental plasticity. An
isolated stem segment, for example, can regenerate
into an entire new plant under appropriate culture
Fig. 12.1 Close-up view of a callus culture.
conditions. For most plant species, some form of
tissue-culture step is therefore a prerequisite for the
successful production of transgenic plants, i.e. plants
carrying the same foreign DNA sequence (the transgene) in every cell. Before considering the various
gene-transfer systems available for plants, we shall
therefore begin with a discussion of the basic principles of plant callus and cell culture. Notably, there
is now increasing interest in the use of whole-plant
(in planta) transformation strategies, in which the
need for tissue culture is minimized or eliminated.
Plant callus and cell culture
Callus culture
Tissue culture is the process whereby small pieces of
living tissue (explants) are isolated from an organism
and grown aseptically for indefinite periods on a
nutrient medium. For successful plant tissue culture,
it is best to start with an explant rich in undetermined cells, e.g. those from the cortex or meristem,
because such cells are capable of rapid proliferation.
The usual explants are buds, root tips, nodal stem
segments or germinating seeds, and these are placed
on suitable culture media where they grow into an
undifferentiated mass known as a callus (Fig. 12.1).
Since the nutrient media used for plants can also
support the growth of microorganisms, the explant
is first washed in a disinfectant such as sodium
hypochlorite or hydrogen peroxide. Once established, the callus can be propagated indefinitely by
subdivision.
For plant cells to develop into a callus, it is essential that the nutrient medium contains the correct
balance of plant hormones (phytohormones), i.e.
auxins, cytokinins and gibberellins, to maintain the
cells in an undifferentiated state (Fig. 12.2). The
absolute amounts required vary for different tissue
explants from different parts of the same plant, and
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CHAPTER 12
HN–CH3
CH2–COOH
N
N
N
N
N
H
H
A cytokinin
(N6-methylaminopurine)
An auxin
(indoleacetic acid (IAA))
OH
CH2
O
C
O
friable calli, and cell lines obtained from these species
are much more homogeneous, allowing either continuous or batchwise cultivation.
If placed in a suitable medium, isolated single cells
from suspension cultures are capable of division.
As with animal cells, conditioned medium may be
necessary for proliferation to occur. Conditioned
medium is prepared by culturing high densities of
cells in fresh medium for a few days and then removing the cells by filter sterilization. Medium conditioned in this way contains essential amino acids,
such as glutamine and serine, as well as growth regulators, such as cytokinins. Provided conditioned
medium is used, single cells can be plated on to solid
media in exactly the same way as microorganisms,
but, instead of forming a colony, plant cells proliferate and form a callus.
HO
CH3
COOH
A gibberellin
Fig. 12.2 The structures of some chemicals which are plant
growth regulators, phytohormones.
for the same explant from different species. Thus
there is no ideal medium. Most of the media in common use consist of inorganic salts, trace metals,
vitamins, organic nitrogen sources (e.g. glycine),
inositol, sucrose and growth regulators. More complex organic nutrients, such as casein hydrolysate,
coconut water or yeast extract, and a gelling agent
are optional extras.
Cell-suspension culture
When callus is transferred into liquid medium and
agitated, the cell mass breaks up to give a suspension
of isolated cells, small clusters of cells and much
larger aggregates. Such suspensions can be maintained indefinitely by subculture but, by virtue of
the presence of aggregates, are extremely heterogeneous. Genetic instability adds to this heterogeneity,
so that long-term culture results in the accumulation of mutations (somaclonal variation), which can
adversely affect the vitality and fertility of regenerated plants. Some species, such as Nicotiana tabacum
(tobacco) and Glycine max (soybean), yield very
Protoplasts
Protoplasts are cells from which the cellulose walls
have been removed. They are very useful for genetic
manipulation because, under certain conditions,
protoplasts from similar or contrasting cell types can
be fused to yield somatic hybrids, a process known as
protoplast fusion. Protoplasts can be produced from
suspension cultures, callus tissue or intact tissues,
e.g. leaves, by mechanical disruption or, preferably,
by treatment with cellulolytic and pectinolytic
enzymes. Pectinase is necessary to break up cell
aggregates into individual cells and the cellulase
digests away the cell wall. After enzyme treatment,
protoplast suspensions are collected by centrifugation, washed in medium without the enzyme, and
separated from intact cells and cell debris by
flotation on a cushion of sucrose. When plated on to
nutrient medium, protoplasts will synthesize new
cell walls within 5–10 days and then initiate cell
division (Fig. 12.3).
Regeneration of fertile plants
The developmental plasticity of plant cells means
that whole fertile plants can often be regenerated
from tissue explants, callus, cell suspensions or protoplasts by placing them on appropriate media. As
discussed above, the maintenance of cells in an
undifferentiated state requires the correct balance
POGC12 9/11/2001 11:14 AM Page 223
Gene transfer to plants
Fig. 12.3 Photomicrograph of tobacco protoplasts.
of phytohormones. However, only cytokinin is
required for shoot culture and only auxin for root
culture, therefore increasing the level of cytokinins
available to the callus induces shoot formation and
increasing the auxin level promotes root formation.
Ultimately, plantlets arise through the development
of adventitious roots on shoot buds or through the
development of shoot buds from tissues formed by
proliferation at the base of rootlets. The formation of
roots and shoots on callus tissue is known as organogenesis. The culture conditions required to achieve
organogenesis vary from species to species, and
have not been determined for every type of callus.
The adventitious organogenesis of shoots and roots
can also occur directly from organized plant tissues,
such as stem segments, without first passing through
a callus stage.
Under certain conditions, cell suspensions or
callus tissue of some plant species can be induced
to undergo a different development process known
as somatic embryogenesis. In this process, the cells
undergo a pattern of differentiation similar to that
seen in zygotes after fertilization, to produce embryoids. These structures are embryo-like but differ from
normal embryos in being produced from somatic
cells and not from the fusion of two germ cells. The
embryoids can develop into fertile plants without the
223
need to induce root and shoot formation on artificial
media. Immature pollen or microspores can also be
induced to form vegetative cells, producing haploid
callus or embryoids. Such cells can be persuaded to
undergo diploidization by treatment with mitotic
inhibitors.
The ease with which plant material is manipulated and interconverted in culture provides many
opportunities for the development of techniques
for gene transfer and the recovery of transgenic
plants (Fig. 12.4). DNA can be introduced into most
types of plant material – protoplasts, cell suspensions, callus, tissue explants, gametes, seeds, zygotes,
embryos, organs and whole plants – so the ability
to recover fertile plants from such material is often
the limiting step in plant genetic engineering rather
than the DNA transfer process itself. It is also possible to maintain transformed plant cell lines or
tissues (e.g. root cultures) producing recombinant
proteins or metabolites, in the same way that cultured animal cells can be used as bioreactors for
valuable products.
Overview of gene-transfer strategies
Gene transfer to plants is achieved using three different methods. The first exploits the natural ability of
certain bacteria of the genus Agrobacterium to naturally transfer DNA to the genomes of infected plant
cells. This generally results in the stable transformation of the infected cell, and the transferred DNA
behaves as a new genetic locus. Initial limitations
with respect to the host range of Agrobacterium
prompted research into alternative methods based
on direct DNA transfer. These include the chemically assisted transformation of protoplasts and the
bombardment of plant material with DNA-coated
microprojectiles. Such strategies can be used for
both transient and stable transformation. Finally,
plant viruses can been used as vectors for gene delivery. The viruses of plants never integrate into the
genome and are not transmitted through seeds, so
stable transformation cannot be achieved. However,
plant viruses often cause systemic infections, resulting in the rapid production of high levels of recombinant protein throughout the plant, and they can be
transmitted through normal infection routes or by
grafting infected scions on to virus-free hosts.
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CHAPTER 12
Agitation in
liquid media
Low auxin +
low cytokinin
Shoots
Protoplasts
Cells
Solid
medium
Callus
Low auxin
+ high
cytokinin
High auxin
+ low
cytokinin
Solid
medium
High
auxin
Plantlets
High
cytokin
Roots
Maintenance
on solid
media with
low auxin
and low
cytokinin
content
Agitation in
liquid media
Callus
Cells
shoots/roots
Explant
on solid
medium
Enzymic digestion
Agitation
in liquid
media
Enzymic digestion
Agrobacterium-mediated
transformation
Crown-gall disease
Crown-gall is a plant tumour that can be induced in
a wide variety of gymnosperms and dicotyledonous
angiosperms (dicots) by inoculation of wound sites
with the Gram-negative soil bacterium A. tumefaciens (Fig. 12.5). The involvement of bacteria in
this disease was established by Smith and Townsend (1907). It was subsequently shown that the
crown-gall tissue represents true oncogenic transformation, since the undifferentiated callus can
be cultivated in vitro even if the bacteria are killed
with antibiotics, and retains its tumorous properties
(Fig. 12.6). These properties include the ability to
form a tumour when grafted on to a healthy plant,
the capacity for unlimited growth as a callus in
tissue culture even in the absence of phytohormones
necessary for the in vitro growth of normal cells,
and the synthesis of opines, such as octopine and
Protoplasts
Fig. 12.4 Summary of the different
cultural manipulations possible with
plant cells, tissues and organs.
nopaline, which are unusual amino acid derivatives
not found in normal plant tissue (Fig. 12.7).
The metabolism of opines is a central feature of
crown-gall disease. Opine synthesis is a property
conferred upon the plant cell when it is colonized by
A. tumefaciens. The type of opine produced is determined not by the host plant but by the bacterial
strain. In general, the bacterium induces the synthesis of an opine that it can catabolize and use as
its sole carbon and nitrogen source. Thus, bacteria
that utilize octopine induce tumours that synthesize
octopine, and those that utilize nopaline induce
tumours that synthesize nopaline (Bomhoff et al.
1976, Montaya et al. 1977).
Tumour-inducing (Ti) plasmids
Since the continued presence of Agrobacterium is not
required to maintain plant cells in their transformed
state, it is clear that some ‘tumour-inducing principle’ is transferred from the bacterium to the plant
at the wound site. Zaenen et al. (1974) first noted
POGC12 9/11/2001 11:14 AM Page 225
Gene transfer to plants
225
Ti plasmids specify the type of opine that is synthesized in the transformed plant tissue and the
opine utilized by the bacteria. Plasmids in the octopine
group are closely related to each other, while those
in the nopaline group are considerably more diverse.
Between the groups, there are four regions of homology, including the genes directly responsible for
tumour formation (Drummond & Chilton 1978,
Engler et al. 1981; Fig. 12.8). It should be noted that
the presence of a plasmid in A. tumefaciens does not
mean that the strain is virulent. Many strains contain very large cryptic plasmids that do not confer
virulence and, in some natural isolates, a cryptic
plasmid is present together with a Ti plasmid.
T-DNA transfer
T-DNA sequences required (in cis) for transfer
Fig. 12.5 Crown gall on blackberry cane. (Photograph
courtesy of Dr C.M.E. Garrett, East Malling Research Station.)
that virulent strains of A. tumefaciens harbour large
plasmids (140–235 kbp), and experiments involving the transfer of such plasmids between various
octopine- and nopaline-utilizing strains soon established that virulence and the ability to use and
induce the synthesis of opines are plasmid-borne
traits. These properties are lost when the bacteria
are cured of their resident plasmid (Van Larbeke
et al. 1974, Watson et al. 1975) but are acquired
by avirulent strains when a virulence plasmid is
reintroduced by conjugation (Bomhoff et al. 1976,
Gordon et al. 1979). The plasmids therefore became
known as tumour-inducing plasmids (Ti plasmids).
Complete Ti-plasmid DNA is not found in plant
tumour cells but a small, specific segment of the
plasmid, about 23 kbp in size, is found integrated in
the plant nuclear DNA at an apparently random
site. This DNA segment is called T-DNA (transferred
DNA) and carries genes that confer both unregulated growth and the ability to synthesize opines
upon the transformed plant tissue. However, these
genes are non-essential for transfer and can be
replaced with foreign DNA (see below). The structure and organization of nopaline plasmid T-DNA
sequences is usually simple, i.e. there is a single
integrated segment. Conversely, octopine T-DNA
comprises two segments: TL (which carries the genes
required for tumour formation) and TR (which carries the genes for opine synthesis). The two segments
are transferred to the plant genome independently
and may be present as multiple copies. The significance of this additional complexity is not clear.
In the Ti plasmid itself, the T-DNA is flanked by
25 bp imperfect direct repeats known as border
sequences, which are conserved between octopine
and nopaline plasmids. The border sequences are
not transferred intact to the plant genome, but they
are involved in the transfer process. The analysis of
junction regions isolated from plant genomic DNA
has shown that the integrated T-DNA end-points lie
internal to the border sequences. The right junction
is rather precise, but the left junction can vary by
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CHAPTER 12
Wound infected with soil
bacterium A. tumefaciens
Crown gall
Excise tissue
from crown gall
Tissue continues to grow
as callus in culture in
absence of added
phytohormones
Carbenicillin in medium
kills Agrobacterium
Callus
Fig. 12.6 Agrobacterium tumefaciens induces plant
tumours.
H
H
R
CO2H
C
NH
CH3
Octopine
R = NH2
CO2H
H
Octopine family
Lysopine
R = NH2(CH2)4
Histopine
R=
(CH2)3
NH
(CH2)3
CH
N
NH
NH
H
C
CO2H
(CH2)2C
Nopaline
R = NH2
C
Nopalinic acid R = NH2(CH2)3
NH
HO2C
NH
Octopinic acid R = NH2(CH2)3
C
R
C
CO2H
H
Nopaline family
Fig. 12.7 Structures of some opines.
POGC12 9/11/2001 11:14 AM Page 227
Gene transfer to plants
227
T-DNA
T-DNA
Virulence
genes
Conjugative
transfer
Octopine
Ti-plasmid
pTiACH5
Virulence
genes
Agrocinopine
catabolism
Conjugative transfer
Fig. 12.8 Ti-plasmid gene maps.
about 100 nucleotides (Yadav et al. 1982, Zambryski
et al. 1982). Deletion of the right border repeat abolishes T-DNA transfer, but the left-hand border surprisingly appears to be non-essential. Experiments
in which the right border repeat alone has been used
have shown that an enhancer, sometimes called the
overdrive sequence, located external to the repeat is
also required for high-efficiency transfer (Shaw et al.
1984, Peralta et al. 1986). The left border repeat has
little transfer activity alone (Jen & Chilton 1986).
Genes required (in trans) for transfer
The genes responsible for T-DNA transfer are located
in a separate part of the Ti plasmid called the vir
(virulence) region. Two of these genes, virA and
virG, are constitutively expressed at a low level and
control the plant-induced activation of the other vir
genes. VirA is a kinase that spans the inner bacterial
membrane and acts as the receptor for certain
phenolic molecules that are released by wounded
plant cells. A large number of such compounds have
been characterized, but one in particular, acetosyringone, has been the most widely used in the laboratory to induce vir gene expression (Stachel et al.
1985; Fig. 12.9). Notably, phenolic compounds such
as acetosyringone do not attract bacteria to wounded
plant cells. Rather, the bacteria appear to respond to
simple molecules, such as sugars and amino acids,
and the vir genes are induced after attachment
(Parke et al. 1987, Loake et al. 1988). Many sugars
also synergize the action of the phenolic signals to
enhance vir gene expression (Shimada et al. 1990).
Activated VirA transphosphorylates the VirG protein, which is a transcriptional activator of the other
vir genes. The VirA and VirG proteins show similar-
Nopaline
Ti-plasmid
pTiC58
Octopine
catabolism
Agropine
catabolism
Nopaline
catabolism
O
O
CH3
C
CH3O
OCH3
OH
Acetosyringone
CH2OH
C
CH3O
OCH3
OH
α-hydroxyacetosyringone
Fig. 12.9 Structures of signal molecules, produced by
wounded plant tissue, which activate T-DNA transfer by
Agrobacterium tumefaciens.
ities to other two-component regulatory systems
common in bacteria (Winans 1992). In addition to
virG, further genes on the bacterial chromosome
also encode transcription factors that regulate vir
gene expression (reviewed by Kado 1998).
The induction of vir gene expression results in the
synthesis of proteins that form a conjugative pilus
through which the T-DNA is transferred to the plant
cell. The components of the pilus are encoded by
genes in the virB operon (reviewed by Lai & Kado
2000). DNA transfer itself is initiated by an endonuclease formed by the products of the virD1 and virD2
genes. This introduces either single-strand nicks or
a double-strand break at the 25 bp borders of the
T-DNA, a process enhanced by the VirC12 and VirC2
proteins, which recognize and bind to the overdrive
enhancer element. The VirD2 protein remains covalently attached to the processed T-DNA. Recent studies
have suggested that the type of T-DNA intermediate
produced (single- or double-stranded) depends on
the type of Ti plasmid, with double-stranded T-DNA
favoured by nopaline plasmids (where the T-DNA is
a single element) and single ‘T strands’ favoured by
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CHAPTER 12
octopine and succinopine plasmids, where the T-DNA
is split into non-contiguous sections (Steck 1997).
T strands are coated with VirE2, a single-stranded
DNA-binding protein. The whole complex, sometimes
dubbed the firecracker complex because of its proposed shape, is then transferred through the pilus
and into the plant cell. It has been proposed that the
VirD2 protein protects the T-DNA against nucleases,
targets the DNA to the plant-cell nucleus and integrates it into the plant genome. The protein has
two distinct nuclear localization signals, with the Cterminal signal thought to play the major role in
targeting the T-DNA (Tinland et al. 1992). It has
been observed that the nucleus of wounded plant cells
often becomes associated with the cytosolic membrane
close to the wound site, suggesting that the T-DNA
could be transferred directly to the nucleus without
extensive exposure to the cytosol (Kahl & Schell
1982). Once in the nucleus, the T-DNA is thought
to integrate through a process of illegitimate recombination, perhaps exploiting naturally occurring
chromosome breaks (Tinland 1996).
The Agrobacterium gene-transfer system appears
to be a highly adapted form of bacterial conjugation.
Many broad-host-range plasmids can transfer from
Agrobacterium to the plant genome using their own
mobilization functions (Buchanan-Wollaston et al.
1987) and the vir genes encode many components
that are common with broad-host-range plasmid
conjugation systems (reviewed by Kado 1998). In
addition to plants, Agrobacterium can transfer DNA
to other bacteria and to yeast. Recently, a novel
insight into the scope of this gene-transfer mechanism was provided by Citovsky and colleagues (Kunik
Gene
et al. 2001) by demonstrating that gene transfer
from Agrobacterium to cultured human cells was also
possible! For the interested reader, T-DNA transfer has been discussed in several comprehensive
reviews (Hooykaas & Schilperoort 1992, Hooykaas
& Beijersbergen 1994, Sheng & Citovsky 1996,
Tzfira et al. 2000, Zupan et al. 2000).
Disarmed Ti-plasmid derivatives
as plant vectors
Function of T-DNA genes
Genetic maps of T-DNA have been obtained by
studying spontaneous and transposon-induced
mutants that affect tumour morphology, generating
tumours that are larger than normal or that show
‘shooty’ or ‘rooty’ phenotypes. Although normal
tumours can grow on medium lacking auxins and
cytokinins, the tumour cells actually contain high
levels of these hormones. Ooms et al. (1981) therefore proposed that the oncogenes carried on the TDNA encoded products involved in phytohormone
synthesis and that the abnormal morphologies of
T-DNA mutants were due to a disturbance in the
balance of plant hormones in the callus. The cloning
and functional analysis of T-DNA genes has confirmed that those with ‘shooty’ mutant phenotypes
encode enzymes for auxin biosynthesis and those
with ‘rooty’ phenotypes are involved in cytokine
production (Weiler & Schroder 1987). Other genes
have been identified as encoding enzymes for opine
synthesis, while the function of some genes remains
unknown (Table 12.1). The transcript maps of
Product
Function
ocs
nos
tms1 (iaaH, auxA)
tms2 (iaaM, auxB )
tmr (ipt, cyt )
tml
Octopine synthase
Nopaline synthase
Tryptophan-2-mono-oxygenase
Indoleacetamide hydrolase
Isopentyl transferase
Unknown
frs
mas
ags
Fructopine synthase
Mannopine synthase
Agropine synthase
Opine synthesis
Opine synthesis
Auxin synthesis
Auxin synthesis
Cytokinin synthesis
Unknown, mutations
affect tumour size
Opine synthesis
Opine synthesis
Opine synthesis
Table 12.1 Functions of some T-DNA
genes in Agrobacterium tumefaciens Ti
plasmids.
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Gene transfer to plants
229
Nopaline T-DNA (23 kb) pTiC58
L border
R border
5
Fig. 12.10 Structure and transcription of
T-DNA. The T-regions of nopaline and
octopine Ti-plasmids have been aligned
to indicate the common DNA sequences.
The size and orientation of each transcript
(numbered) is indicated by arrows. Genetic
loci, defined by deletion and transposon
mutagenesis, are shown as follows: nos,
nopaline synthase; ocs, octopine synthase;
tms, shooty tumour; tmr, rooty tumour.
2
1
tms
4 6a6b 3
tmr tml nos
TR-DNA
Octopine TL-DNA (13.6 kb) pTiAch5
L border
T-DNAs from a nopaline plasmid (pTiC58) and an
octopine plasmid (pTiAch5) are shown in Fig. 12.10
(Willmitzer et al. 1982, 1983, Winter et al. 1984).
Interestingly, nucleotide sequencing has revealed
that the T-DNA genes have promoter elements and
polyadenylation sites that are eukaryotic in nature
(De Greve et al. 1982b, Depicker et al. 1982, Bevan
et al. 1983a). This explains how genes from a bacterial
plasmid come to be expressed when transferred to
the plant nucleus. It is possible that the sequences
may have been captured from plants during the evolution of the Ti plasmid. The ability of Agrobacterium
to induce tumours in a wide variety of plants suggested that T-DNA promoters, such as those of the
ocs (octopine synthase) and nos (nopaline synthase)
genes, could be useful to drive transgene expression. These and other promoters used for transgene
expression in plants are discussed in Box 12.1.
R border
7
5
2
1
tms
L border
R border
4 6a6b 3
4’ 3’ 2’
tmr tml ocs
frs
1’
0’
mas ags
resulting construct was called pGV3850 (Fig. 12.11).
Agrobacterium carrying this plasmid transferred
the modified T-DNA to plant cells. As expected, no
tumour cells were produced, but the fact that transfer had taken place was evident when the cells were
screened for nopaline production and found to be
positive. Callus tissue could be cultured from these
nopaline-positive cells if suitable phytohormones
were provided, and fertile adult plants were regenerated by hormone induction of plantlets.
The creation of disarmed T-DNA was an important step forward, but the absence of tumour formation made it necessary to use an alternative method
to identify transformed plant cells. In the experiment
described above, opine production was exploited
as a screenable phenotype, and the ocs and nos
genes have been widely used as screenable markers
pBR322
Prototype disarmed Ti vectors
We have seen that the Ti plasmid is a natural vector
for genetically engineering plant cells because it can
transfer its T-DNA from the bacterium to the plant
genome. However, wild-type Ti plasmids are not
suitable as general gene vectors because the T-DNA
contains oncogenes that cause disorganized growth
of the recipient plant cells. To be able to regenerate plants efficiently, we must use vectors in which
the T-DNA has been disarmed by making it nononcogenic. This is most effectively achieved simply by
deleting all of its oncogenes. For example, Zambryski
et al. (1983) substituted pBR322 sequences for
almost all of the T-DNA of pTiC58, leaving only the
left and right border regions and the nos gene. The
L border
AmpR
nos
R border
pGV3850
(PTiC58 derivative)
vir genes
Fig. 12.11 Structure of the Ti-plasmid pGV3850, in which
the T-DNA has been disarmed.
POGC12 9/11/2001 11:14 AM Page 230
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CHAPTER 12
Box 12.1 Control of transgene expression in plants
Promoters
To achieve high-level and constitutive transgene
expression in plants, a very active promoter
is required. In dicots, promoters from the
Agrobacterium nopaline synthase (nos), octopine
synthase (ocs) and mannopine synthase (mas)
genes have been widely used. These are constitutive
and also moderately induced by wounding
(An et al. 1990, Langridge et al. 1989). The most
popular promoter for transgene expression in
dicots is the 35S RNA promoter from cauliflower
mosaic virus (CaMV 35S). This is very active, but can
be improved still further by duplicating the enhancer
region (Rathus et al. 1993). These promoters have a
much lower activity in monocots, and duplicating
the CaMV 35S enhancer has little effect. Alternative
promoters have therefore been sought for transgene
expression in cereals (reviewed by McElroy & Brettel
1994). The rice actin-1 and maize ubiquitin-1
promoters have been widely used for this purpose
(McElroy et al. 1995, Christensen & Quail 1996).
As well as constitutive promoters, a large number
of promoters have been used to direct transgene
expression in particular tissues. In monocots,
promoters from seed storage-protein genes, such as
maize zein, wheat glutenin and rice glutelins, have
been used to target transgene expression to the
seeds, which is beneficial for the accumulation of
recombinant proteins (Wu et al. 1998; reviewed by
Bilan et al. 1999). Promoters targeting transgene
expression to green tissue are also useful (e.g.
Graham et al. 1997, Datta et al. 1998). We discuss
inducible expression systems for animals and plants
in Chapter 13.
Other components of the expression vector
As discussed for animal cells (Box 10.2), other
sequences in the expression vector also influence
transgene expression. Generally, the presence of
an intron in a plant expression cassette increases
the activity of the promoter (Bilan et al. 1999).
The insertion of a heterologous intron enhances the
activity of the CaMV 35S promoter in monocots
(e.g. see Mascarenhas et al. 1990, Vain et al. 1996)
and constructs containing the actin or ubiquitin
promoters generally include the first intron of the
gene (McElroy et al. 1991). All transgenes must
include a polyadenylation site, which in most cases
is derived from the Agrobacterium nos gene or
the CaMV 35S RNA. Whereas in animals it is
conventional to remove untranslated regions
from the expression construct, a number of such
sequences in plants have been identified as
translational enhancers. For example, the 5′ leader
sequence of the tobacco mosaic virus RNA, known
as the omega sequence, can increase transgene
expression up to 80-fold (reviewed by Futterer
& Hohn 1996, Gallie 1996). As in animals, the
translational start site should conform to Kozak’s
consensus (Kozak 1999; see Box 10.2 for details)
and the transgene should be codon-optimized for
the expression host. A good example of the latter is
the use of codon-optimized insecticidal toxin genes
from Bacillus thuringiensis for expression in
transgenic crops, leading to dramatically increased
expression levels compared with the unmodified
genes (Koziel et al. 1996). Also, the inclusion of
targeting information in the expression cassette may
be beneficial for the accumulation of recombinant
proteins. For example, recombinant antibodies
expressed in plants are much more stable if targeted
to the endoplasmic reticulum (ER), since this provides
a favourable molecular environment for folding and
assembly. Targeting is achieved using an N-terminal
signal sequence to direct the ribosome to the ER
and a C-terminal tetrapeptide retrieval signal,
KDEL, which causes accumulation in the ER lumen
(Horvath et al. 2000).
POGC12 9/11/2001 11:14 AM Page 231
Gene transfer to plants
(reviewed by Dessaux & Petit 1994). However, there
are several drawbacks associated with this system,
particularly the necessity to carry out enzymatic
assays on all potential transformants. To provide a
more convenient way to identify transformed plant
231
cells, dominant selectable markers have been inserted
into the T-DNA, so that transformed plant cells
can be selected on the basis of drug or herbicide
resistance. The use of selectable markers in plants
is discussed in more detail in Box 12.2.
Box 12.2 Selectable markers for plants
Until recently, almost all selectable markers used in
plants were dominant selectable markers, providing
resistance to either antibiotics or herbicides (see
table). Some plants, particularly monocots, are
naturally tolerant of kanamycin, and this antibiotic
may also interfere with regeneration. In these species,
alternative systems, such as hygromycin or
phosphinothricin selection, are preferred.
The introduction of markers such as nptII, hpt
and dhfr into the T-DNA of disarmed Ti plasmids
provided the first convenient methods to identify
transformed plant tissue, and hence opened the way
for Agrobacterium to be used as a general plant
transformation system. The marker and experimental
transgene can be cloned in tandem on the same
T-DNA. In such cases, it is better for the selectable
marker to be placed adjacent to the left border
repeat, since this is transferred to the plant last
(Sheng & Citovsky 1996). This strategy reduces the
likelihood of obtaining plants under selection
Marker
Drug resistance markers
aad (preferred for chloroplast
transformation (see p. 240)
ble
dhfr
hpt
nptII and aphII
gat
Herbicide resistance markers
bar and pat
csr1-1
dhps (sul)
epsp
containing the marker alone and not the transgene
of interest (see Hellens et al. 2000a). Alternatively,
cotransformation can be achieved using
Agrobacterium strains containing two plasmids or by
co-inoculating plants with different Agrobacterium
strains, each containing a single plasmid. Although
there is some controversy surrounding the fate of
co-introduced T-DNA sequences, it appears that
nopaline-type plasmids favour the co-integration
of multiple T-DNAs at the same locus, often in
an inverted repeat pattern, while octopine-type
plasmids favour independent integration sites,
which can segregate in progeny plants (Depicker
et al. 1985, Jones et al. 1987, Jorgensen et al.
1987, De Block & Debrouwer 1991). In direct
transformation methods, the selectable marker
can be included either on the same vector as the
experimental transgene or on a separate vector,
since cotransformation occurs at a high frequency
(Schocher et al. 1986, Christou & Swain 1990).
Selection
Trimethoprim, streptomycin,
spectinomycin, sulphonamides
Bleomycin
Methotrexate
Hygromycin
Kanamycin, neomycin, G418
Gentamicin
Phosphinothricin (bialaphos,
glufosinate ammonium, Basta)
Chlorsulphuron
Sulphonamides (Asualam)
Glyphosate
References
Svab et al. 1990a
Hille et al. 1986
Eichholtz et al. 1987
Van den Elzen et al. 1985
Pridmore 1987
Hayford et al. 1988
De Block et al. 1987
Haughn et al. 1988
Guerineau & Mullineaux 1989
Shah et al. 1986
continued
POGC12 9/11/2001 11:14 AM Page 232
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CHAPTER 12
Box 12.2 continued
Recently, public concern that antibiotic- and
herbicide-resistance markers could pose a threat to
health or the environment has prompted research
into alternative innocuous marker systems. One
example is the E. coli manA gene, which encodes
mannose phosphate isomerase and confers upon
transformed cells the ability to use mannose as a
sole carbon source (Negrotto et al. 2000). Another
is the A. tumefaciens ipt (isopentyl transferase)
gene, located on the T-DNA, which induces
cytokinin synthesis and can be used to select
plants on the basis of their ability to produce
shoots from callus on medium lacking cytokinins
(Kunkel et al. 1999). Other strategies have also
been explored, such as eliminating markers by
sexual crossing (Komari et al. 1996), transposition
(Goldsbrough et al. 1993) or site-specific
Cointegrate vectors
Although disarmed derivatives of wild-type Ti plasmids can be used for plant transformation, they are
not particularly convenient as experimental gene
vectors, because their large size makes them difficult to manipulate in vitro and there are no unique
restriction sites in the T-DNA. Initially, this problem
was addressed by the construction of cointegrate
vectors. T-DNA isolated from a parent Ti plasmid
was subcloned in a conventional Escherichia coli
plasmid vector for easy manipulation, producing
a so-called intermediate vector (Matzke & Chilton
1981). These vectors were incapable of replication
in A. tumefaciens and also lacked conjugation functions. Transfer was achieved using a ‘triparental
mating’ in which three bacterial strains were mixed
together: (i) an E. coli strain carrying a helper
plasmid able to mobilize the intermediate vector in
trans; (ii) the E. coli strain carrying the recombinant
intermediate vector; and (iii) A. tumefaciens carrying
the Ti plasmid. Conjugation between the two E. coli
strains transferred the helper plasmid to the carrier
of the intermediate vector, which was in turn mobilized and transferred to the recipient Agrobacterium.
recombination (Dale & Ow 1991, Russel et al.
1992, Zubko et al. 2000). We consider the use
of Cre-loxP for marker excision in transgenic
organisms in Chapter 13. To verify the successful
elimination of particular genes, counterselectable
markers are required (e.g. see the discussion of Tk
as a couterselectable marker for gene targeting in
mice, p. 206). In plants, the A. tumefaciens T-DNA
gene tms2 has been used as a negative marker.
This encodes indoleacetamide hydrolase, an enzyme
that converts naphthaleneacetamide (NAM) into
the potent auxin naphthaleneacetic acid. In the
presence of exogenously applied NAM, transformed
callus is unable to produce shoots due to the excess
levels of auxin, therefore only callus lacking the
gene is able to regenerate into full transgenic plants
(Sundaresan et al. 1995).
Homologous recombination between the T-DNA
sequences of the Ti plasmid and intermediate vector
then resulted in the formation of a large cointegrate
plasmid, from which the recombinant T-DNA was
transferred to the plant genome (Fig. 12.12). In
the cointegrate vector system, maintenance of the
recombinant T-DNA is dependent on recombination, which is enhanced if there is an extensive
homology region shared by the two plasmids, as in
Ti plasmid pGV3850, which carries a segment of
the pBR322 backbone in its T-DNA.
Binary vectors
Although intermediate vectors have been widely
used, the large cointegrates are not necessary for
transformation. The vir genes of the Ti plasmid function in trans and can act on any T-DNA sequence
present in the same cell. Therefore, the vir genes
and the disarmed T-DNA containing the transgene
can be supplied on separate plasmids, and this is the
principle of binary vector systems (Hoekma et al.
1983, Bevan 1984). The T-DNA can be subcloned
on a small E. coli plasmid for ease of manipulation.
This plasmid, called mini-Ti or micro-Ti, can be
POGC12 9/11/2001 11:14 AM Page 233
Gene transfer to plants
233
Intermediate vector transferred
into Agrobacterium by
conjugation. Unable to replicate
autonomously in Agrobacterium
Disarmed Ti-vector pGV3850
resident in Agrobacterium
neo
nos
KanR
pGV3850
AmpR
pBR322 pBR322
AmpR
vir genes
Foreign gene
inserted
Homologous recombination between DNA regions derived from pBR322
Cointegrate formation
Select for maintenance of
cointegrate by kanamycin
Foreign gene
neo
AmpR
KanR
nos
AmpR
vir genes
Fig. 12.12 Production of recombinant
disarmed Ti plasmid by cointegrate
formation.
Cointegrate disarmed Ti-plasmid
introduced into an Agrobacterium strain carrying
a Ti plasmid from which the T-DNA has been
removed. The vir functions are supplied in trans,
causing transfer of the recombinant T-DNA to the
plant genome. The T-DNA plasmid can be introduced into Agrobacterium by triparental matings or
by a more simple transformation procedure, such as
electroporation (Cangelosi et al. 1991).
Most contemporary Ti-plasmid transformation
systems are based on a binary principle, in which
the T-DNA is maintained on a shuttle vector with
a broad-host-range origin of replication, such as
RK2 (which functions in both A. tumefaciens and
E. coli), or separate origins for each species. An
independently replicating vector is advantageous
because maintenance of the T-DNA is not reliant
POGC12 9/11/2001 11:14 AM Page 234
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CHAPTER 12
Bsp HI (3031)
pSa-ORI
Bgl II (536)
Hpa I (568)
Asp 7181 (783)
Apa I (793)
Xho I (798)
Sal I (804)
Cla I (814)
Eco RV (827)
Eco RI (831)
Pst I (841)
Sma I (845)
Bam HI (849)
Spe I (855)
Xba I (861)
Not I (868)
Sac II (880)
Sac I (889)
LB
nptI
lacZ
pGreenII 0000
3304 bp
RB
Bsp HI (2070)
ColEI ori
Stu I (1292)
Bgl II (1345)
on recombination, and the binary vector’s copy
number is not determined by the Ti plasmid, making
the identification of transformants much easier.
All the conveniences of bacterial cloning plasmids
have been incorporated into binary vectors, such as
multiple unique restriction sites in the T-DNA region
to facilitate subcloning, the lacZ gene for blue–white
screening (McBride & Summerfelt 1990) and a λ cos
site for preparing cosmid libraries (Lazo et al. 1991,
Ma et al. 1992). A current binary vector, pGreen,
is shown in Fig. 12.13 (Hellens et al. 2000b). This
plasmid is less than 5 kbp in size and has 18 unique
restriction sites in the T-DNA, because the T-DNA is
entirely synthetic. It has a lacZ gene for blue–white
selection of recombinants, and a selectable marker
that can be used both in bacteria and in the transformed plants. The progressive reduction in size has
been made possible by removing essential genes
required for replication in Agrobacterium and transferring these genes to the bacterium’s genome or
on to a helper plasmid. The pGreen plasmid, for
example, contains the Sa origin of replication, which
is much smaller than the more traditional Ri and
RK2 regions. Furthermore, an essential replicase gene
is housed on a second plasmid, called pSoup, resident
within the bacterium. All conjugation functions
have also been removed, so this plasmid can only
be introduced into Agrobacterium by transformation
(Hellens et al. 2000b).
Fig. 12.13 The small and versatile
binary vector pGreen, reproduced with
permission of Roger Hellens and Phil
Mullineaux.
A simple experimental procedure for
Agrobacterium-mediated
transformation
Once the principle of selectable, disarmed T-DNA
vectors was established, there followed an explosion
in the number of experiments involving DNA transfer to plants. Variations on the simple general
protocol of Horsch et al. (1985) have been widely
used for dicot plants (Fig. 12.14). In the original
report, small discs (a few millimetres in diameter)
were punched from leaves, surface-sterilized and
inoculated in a medium containing A. tumefaciens
transformed with the recombinant disarmed T-DNA
(as a cointegrate or binary vector). The foreign DNA
contained a chimeric neo gene conferring resistance
to the antibiotic kanamycin. The discs were cultured
for 2 days and transferred to medium containing
kanamycin, to select for the transferred neo gene,
and carbenicillin, to kill the Agrobacterium. After
2–4 weeks, developing shoots were excised from the
callus and transplanted to root-induction medium.
Rooted plantlets were subsequently transplanted
to soil, about 4–7 weeks after the inoculation step.
This method has the advantage of being simple
and relatively rapid. It is superior to previous
methods, in which transformed plants were regenerated from protoplast-derived callus, the protoplasts
having been transformed by cocultivation with the
POGC12 9/11/2001 11:14 AM Page 235
Gene transfer to plants
235
Surface-sterilize
Inoculation
Culture overnight
in liquid with
Agrobacterium to
infect cut edges of disc
Small discs
punched
from leaf
Blot disc
dry
Leaf disc
Feeder plate
Filter paper
Shoot-inducing solid
medium + kanamycin
+ carbenicillin
Culture for about
20 days
A layer of feeder
cells, previously
grown in
suspension
Shoot-inducing
solid medium
(high in cytokinin)
Culture for 2 days
Shoot
formation
Excise
shooted callus,
transfer to
root-inducing
medium
Fig. 12.14 Leaf-disc transformation by
Agrobacterium tumefaciens.
Agrobacterium (De Block et al. 1984, Horsch et al.
1984). Contemporary protocols for the Agrobacteriummediated transformation of many solanaceous
plants are variations on the theme of the leaf-disc
protocol, although the optimal explant must be
determined for each species. Alternative procedures
are required for the transformation of monocots, as
discussed below.
Agrobacterium and monocots
Until recently, most monocotyledonous plants (monocots) were thought to be outside the host range of
Agrobacterium, prompting research into alternative
transformation methods, as discussed below. During
the 1980s, limited evidence accumulated showing
Root-inducing solid
medium (high in
auxin) + kanamycin +
carbenicillin
Transfer plantlets to
soil as soon as roots
appear 4–7 weeks
after inoculation
that some monocots might be susceptible to Agrobacterium infection (see, for example, the discussion
of agroinfection with maize streak virus DNA on
p. 243). However, in most cases there was no
convincing evidence for T-DNA integration into the
plant genome. In the laboratory, it proved possible
to induce tumours in certain monocot species, such
as asparagus (Hernalsteens et al. 1984) and yam
(Schafer et al. 1987). In the latter case, an important
factor in the success of the experiment was pretreatment of the Agrobacterium suspension with wound
exudate from potato tubers. It has been argued that
Agrobacterium infection of monocots is inefficient
because wounded monocot tissues do not produce
phenolics, such as acetosyringone, at sufficient
levels to induce vir gene expression.
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CHAPTER 12
In the last 10 years, amazing progress has
been made in the transformation of cereals using
Agrobacterium. The first species to be transformed
was rice. The nptII gene was used as a selectable
marker, and successful transformation was demonstrated both by the resistance of transgenic callus to
kanamycin or G418 and by the presence of T-DNA
in the genome (Raineri et al. 1990, Chan et al. 1992,
1993). However, these antibiotics interfere with the
regeneration of rice plants, so only four transgenic
plants were produced in these early studies. The use
of an alternative marker conferring resistance to
hygromycin allowed the regeneration of large numbers of transgenic japonica rice plants (Hiei et al.
1994), and the same selection strategy has been
used to produce transgenic rice plants representing the other important subspecies, indica and
javanica (Dong et al. 1996, Rashid et al. 1996).
More recently, efficient Agrobacterium-mediated
transformation has become possible for other important cereals, including maize (Ishida et al. 1996),
wheat (Cheng et al. 1997), barley (Tingay et al. 1997)
and sugar cane (Arencibia et al. 1998).
The breakthrough in cereal transformation using
Agrobacterium reflected the recognition of a number
of key factors required for efficient infection and gene
transfer to monocots. The use of explants containing
a high proportion of actively dividing cells, such
as embryos or apical meristems, was found to
greatly increase transformation efficiency, probably
because DNA synthesis and cell division favour the
integration of exogenous DNA. In dicots, cell division is often induced by wounding, whereas wound
sites in monocots tend to become lignified. This probably explains why traditional procedures, such as
the leaf-disc method, are inefficient in monocots.
Hiei et al. (1994) showed that the cocultivation of
Agrobacterium and rice embryos in the presence
of 100 mmol/l acetosyringone was a critical factor
for successful transformation. Transformation efficiency is increased further by the use of vectors with
enhanced virulence functions. The modification of
Agrobacterium for increased virulence has been
achieved by increasing the expression of virG (which
in turn boosts the expression of the other vir genes)
and/or the expression of virE1, which is a major limiting factor in T-DNA transfer (reviewed by Sheng
& Citovsky 1996), resulting in so-called supervirulent
bacterial strains, such as AGL-1. Komari et al. (1996)
used a different strategy, in which a portion of the
virulence region from the Ti plasmid of supervirulent
strain A281 was transferred to the T-DNA-carrying
plasmid to generate a so-called ‘superbinary vector’.
The advantage of the latter technique is that the
superbinary vector can be used in any Agrobacterium
strain.
High-capacity binary vectors
A precise upper limit for T-DNA transfer has not
been established. It is greater than 50 kb (HerreraEstrella et al. 1983a), but using standard vectors
it is difficult to routinely transfer inserts larger
than 30 kb due to instability in the bacterial host.
The analysis of very large genes or the transfer of
multiple genes (such as those encoding sequentially
acting enzymes of a metabolic pathway) can now
be achieved thanks to the development of highcapacity binary vectors based on the artificialchromosome-type vectors used in E. coli. The first to
be described was BIBAC2 (Hamilton 1997). This
contains an F-plasmid origin of replication and
is modelled on the bacterial artificial chromosome
(BAC) (p. 67). The basic vector transforms tobacco
with high efficiency, but the efficiency of transformation drops substantially when large inserts are used.
This vector has been used to introduce 150 kbp of
human DNA flanked by T-DNA borders into the
tobacco genome, although virulence helper plasmids
supplying high levels of VirG and VirE in trans were
critical for successful DNA transfer (Hamilton et al.
1996). An alternative vector carrying a P1 origin
of replication and modelled on the P1 artificial
chromosome (PAC) (p. 68) was constructed by
Liu et al. (1999). This transformation-competent
bacterial artificial chromosome (TAC) vector was
used to introduce up to 80 kbp of genomic DNA
into Arabidopsis and, while there was some loss of
efficiency with the larger inserts, it was still possible
to produce many transgenic plants. Both vectors contain a kanamycin-resistance marker for selection in
bacteria and hpt for hygromycin selection in transgenic plants. For the reasons discussed in Box 12.2,
the hpt marker gene is placed adjacent to the right
border T-DNA repeat. Both vectors also contain the
Ri origin for maintenance in Agrobacterium and,
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Gene transfer to plants
within the T-DNA region, the sacB marker for
negative selection in E. coli, interrupted by a polylinker for cloning foreign DNA.
One of the most attractive uses of high-capacity
binary vectors is for the positional cloning of genes
identified by mutation. The ability to introduce large
segments of DNA into the plant genome effectively bridges the gap between genetic mapping and
sequencing, allowing the position of mutant genes
to be narrowed down by complementation. Genomic
libraries have been established for several plant
species in BIBAC2 and TAC vectors (Hamilton et al.
1999, Shibata & Lui 2000) and a number of novel
genes have been isolated (e.g. Sawa et al. 1999,
Kubo & Kakimoto 2001).
Agrobacterium rhizogenes and
Ri plasmids
Agrobacterium rhizogenes causes hairy-root disease
in plants and this is induced by root-inducing (Ri)
plasmids, which are analogous to the Ti plasmids of
A. tumefaciens. The Ri T-DNA includes genes homologous to the iaaM (tryptophan 2-mono-oxygenase)
and iaaH (indoleacetamide hydrolase) genes of A.
tumefaciens. Four other genes present in the Ri T-DNA
are named rol for root locus. Two of these, rolB and
rolC, encode P-glucosidases able to hydrolyse indoleand cytokine-N-glucosides. A. rhizogenes therefore
appears to alter plant physiology by releasing free
hormones from inactive or less active conjugated
forms (Estruch et al. 1991a,b).
Ri plasmids are of interest from the point of view of
vector development, because opine-producing root
tissue induced by Ri plasmids in a variety of dicots
can be regenerated into whole plants by manipulation of phytohormones in the culture medium. Ri TDNA is transmitted sexually by these plants and
affects a variety of morphological and physiological
traits, but does not in general appear deleterious.
The Ri plasmids therefore appear to be already
equivalent to disarmed Ti plasmids (Tepfer 1984).
Transformed roots can also be maintained as hairyroot cultures, which have the potential to produce
certain valuable secondary metabolites at higher
levels than suspension cultures and are much more
genetically stable (Hamil et al. 1987, Signs & Flores
1990). The major limitation for the commercial use
237
of hairy-root cultures is the difficulty involved in
scale-up, since each culture comprises a heterogeneous mass of interconnected tissue, with highly
uneven distribution (reviewed by Giri & Narassu
2000).
Many of the principles explained in the context of
disarmed Ti plasmids are applicable to Ri plasmids.
A cointegrate vector system has been developed
(Jensen et al. 1986) and applied to the study of
nodulation in transgenic legumes. Van Sluys et al.
(1987) have exploited the fact that Agrobacterium
containing both an Ri plasmid and a disarmed Ti
plasmid can frequently cotransfer both plasmids. The
Ri plasmid induces hairy-root disease in recipient
Arabidopsis and carrot cells, serving as a transformation marker for the cotransferred recombinant TDNA and allowing regeneration of intact plants. No
drug-resistance marker on the T-DNA is necessary
with this plasmid combination.
Direct DNA transfer to plants
Protoplast transformation
Until comparatively recently, the limited host range
of A. tumefaciens precluded its use for the genetic
manipulation of a large number of plant species,
including most monocots. At first, the only alternative to Agrobacterium-mediated transformation was
the introduction of DNA into protoplasts. This process has much in common with the transfection
of animal cells (Chapter 10). The protoplasts must
initially be persuaded to take up DNA from their
surroundings, after which the DNA integrates stably
into the genome in a proportion of these transfected
cells. Gene transfer across the protoplast membrane
is promoted by a number of chemicals, of which
polyethylene glycol has become the most widely
used, due to the availability of simple transformation
protocols (Negrutiu et al. 1987). Alternatively, DNA
uptake may be induced by electroporation, which
has also become a favoured technique (Shillito et al.
1985). As with animal cells, the introduction of a
selectable marker gene along with the transgene of
interest is required for the identification of stable
transformants. This can be achieved using plasmid
vectors carrying both the marker and the transgene
of interest, but the use of separate vectors also results
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CHAPTER 12
in a high frequency of cotransformation (Schocher
et al. 1986). Putative transformants are transferred
to selective medium, where surviving protoplasts
regenerate their cell walls and commence cell division, producing a callus. Subsequent manipulation
of the culture conditions then makes it possible to
induce shoot and root development, culminating in
the recovery of fertile transgenic plants. The major
limitation of protoplast transformation is not the
gene-transfer process itself, but the ability of the host
species to regenerate from protoplasts. A general
observation is that dicots are more amenable than
monocots to this process. In species where regeneration is possible, an advantage of the technique is that
protoplasts can be cryopreserved and retain their
regenerative potential (DiMaio & Shillito 1989).
The first transformation experiments concentrated
on species such as tobacco and petunia in which
protoplast-to-plant regeneration is well documented.
An early example is provided by Meyer et al. (1987),
who constructed a plasmid vector containing the
nptII marker gene, and a maize complementary DNA
(cDNA) encoding the enzyme dihydroquercetin 4reductase, which is involved in anthocyanin pigment biosynthesis. The transgene was driven by the
strong and constitutive cauliflower mosaic virus
(CaMV) 35S promoter. Protoplasts of a mutant,
white-coloured petunia strain were transformed
with the recombinant plasmid by electroporation
and then selected on kanamycin-supplemented
medium. After a few days, surviving protoplasts had
given rise to microcalli, which could be induced to
regenerate into whole plants. The flowers produced
by these plants were brick-red instead of white,
showing that the maize cDNA had integrated into
the genome and was expressed.
After successful experiments using model dicots,
protoplast transformation was attempted in monocots, for which no alternative gene-transfer system
was then available. In the first such experiments,
involving wheat (Lorz et al. 1985) and the Italian
ryegrass Lolium multiflorum (Potrykus et al. 1985b),
protoplast transformation was achieved and transgenic callus obtained, but it was not possible to
recover transgenic plants. The inability of most
monocots to regenerate from protoplasts may reflect
the loss of competence to respond to tissue-culture
conditions as the cells differentiate. In cereals and
grasses, this has been addressed to a certain extent
by using embryogenic suspension cultures as a source
of protoplasts. Additionally, since many monocot
species are naturally tolerant to kanamycin, the
nptII marker used in the initial experiments was
replaced with alternative markers conferring resistance to hygromycin or phosphinothricin. With these
modifications, it has been possible to regenerate
transgenic plants representing certain varieties of rice
and maize with reasonable efficiency (Shimamoto
et al. 1988, Datta et al. 1990, Omirulleh et al. 1993).
However, the extended tissue-culture step is unfavourable, often resulting in sterility and other phenotypic
abnormalities in the regenerated plants. The transformation of protoplasts derived from stomatal guard
cells has recently been identified as an efficient and
genotype-independent method for the production of
transgenic sugar beet (Hall et al. 1996).
Particle bombardment
An alternative procedure for plant transformation
was introduced in 1987, involving the use of a
modified shotgun to accelerate small (1–4 µm) metal
particles into plant cells at a velocity sufficient to
penetrate the cell wall (~250 m/s). In the initial
test system, intact onion epidermis was bombarded
with tungsten particles coated in tobacco mosaic
virus (TMV) RNA. Three days after bombardment,
approximately 40% of the onion cells that contained
particles also showed evidence of TMV replication
(Sanford et al. 1987). A plasmid containing the cat
reporter gene driven by the CaMV 35S promoter
was then tested to determine whether DNA could
be delivered by the same method. Analysis of the
epidermal tissue 3 days after bombardment revealed
high levels of transient chloramphenicol transacetylase (CAT) activity (Klein et al. 1987).
The stable transformation of explants from
several plant species was achieved soon after these
initial experiments. Early reports included the transformation of soybean (Christou et al. 1988), tobacco
(Klein et al. 1988b) and maize (Klein et al. 1988a). In
each case, the nptII gene was used as a selectable
marker and transformation was confirmed by the
survival of callus tissue on kanamycin-supplemented
medium. The ability to stably transform plant cells
by this method offered the exciting possibility of
POGC12 9/11/2001 11:14 AM Page 239
Gene transfer to plants
generating transgenic plants representing species
that were, at the time, intractable to other transformation procedures. In the first such report, transgenic soybean plants were produced from meristem
tissue isolated from immature seeds (McCabe et al.
1988). In this experiment, the screenable marker
gene gusA was introduced by particle bombardment
and transgenic plants were recovered in the absence
of selection by screening for β-glucuronidase (GUS)
activity (Box 13.1). Other early successes included
cotton, papaya, maize and tobacco (Finer & McMullen
1990, Fitch et al. 1990, Fromm et al. 1990, GordonKamm et al. 1990, Tomes et al. 1990).
There appears to be no intrinsic limitation to
the scope of this procedure, since DNA delivery is
governed entirely by physical parameters. Many
different types of plant material have been used
as transformation targets, including callus, cellsuspension cultures and organized tissues, such as
immature embryos, meristems and leaves. The
number of species in which transgenic plants can be
produced using variants of particle bombardment
has therefore increased dramatically over the last
10 years. Notable successes include almost all of the
commercially important cereals, i.e. rice (Christou
et al. 1991), wheat (Vasil et al. 1992), oat (Somers
et al. 1992, Torbert et al. 1995), sugar cane (Bower
& Birch 1992) and barley (Wan & Lemaux 1994,
Hagio et al. 1995).
The original gunpowder-driven device has been
improved and modified, resulting in greater control
over particle velocity and hence greater reproducibility of transformation conditions. An apparatus based on electric discharge (McCabe & Christou
1993) has been particularly useful for the development of variety-independent gene-transfer methods
for the more recalcitrant cereals and legumes. Several
instruments have been developed where particle
acceleration is controlled by pressurized gas. These
include a pneumatic apparatus (Iida et al. 1990), a
‘particle inflow gun’ using flowing helium (Finer
et al. 1992, Takeuchi et al. 1992) and a device
utilizing compressed helium (Sanford et al. 1991).
Physical parameters, such as particle size and acceleration (which affect the depth of penetration and
the amount of tissue damage), as well as the amount
and conformation of the DNA used to coat the particles, must be optimized for each species and type of
239
explant. However, the nature of the transformation
target is probably the most important single variable
in the success of gene transfer. The pretreatment of
explants with an osmoticum has often been shown
to improve transformation efficiency, probably by
preventing the deflection of particles by films or
droplets of water. Factors influencing the success of
gene transfer by particle bombardment have been
extensively reviewed (Sanford et al. 1993, Birch &
Bower 1994).
Other direct DNA-transfer methods
There is a great diversity of approaches for gene
transfer to animal cells and many of the same methods have been attempted in plants. Electroporation
has been used to transform not only protoplasts (see
above) but also walled plant cells, either growing in
suspension or as part of intact tissues. In many cases,
the target cells have been wounded or pretreated
with enzymes in order to facilitate gene transfer (e.g.
D’Halluin et al. 1992, Laursen et al. 1994). It has
been shown, however, that immature rice, wheat
and maize embryos can be transformed using electroporation without any form of pretreatment (Kloti
et al. 1993, Xu & Li 1994). Other transformation
methods also involve perforation of the cell, including
the use of silicon carbide whiskers (Thompson et al.
1995, Nagatani et al. 1997), ultrasound (Zhang et al.
1991) or a finely focused laser beam (Hoffman
1996). In most of these cases, only transient expression of the introduced DNA has been achieved,
although transgenic maize plants have been recovered following whisker-mediated transformation.
Finally, microinjection has been used to introduce
DNA directly into the fertilized eggs of many animals
(Chapter 10). In plants, microinjection of DNA into
zygotes may also be the most direct way to produce
transgenics, but so far the technique is inefficient and
not widely used (Leduc et al. 1996, Holm et al. 2000).
In planta transformation
Until recently, gene transfer to plants involved the
use of cells or explants as transformation targets and
an obligatory tissue-culture step was needed for the
regeneration of whole fertile plants. Experiments
using the model dicot Arabidopsis thaliana have led
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CHAPTER 12
the way in the development of so-called in planta
transformation techniques, where the need for
tissue culture is minimized or eliminated altogether.
Such methods involve the introduction of DNA,
either by Agrobacterium or by direct transfer, into
intact plants. The procedure is carried out at an
appropriate time in the plant’s life cycle, so that
the DNA becomes incorporated into cells that will
contribute to the germ line, directly into the germ
cells themselves (often at around the time of fertilization) or into the very early plant embryo. Generally,
in planta transformation methods have a very low
efficiency, so the small size of Arabidopsis and its
ability to produce over 10 000 seeds per plant is
advantageous. This limitation has so far prevented
in planta techniques from being widely adopted for
other plant species.
The first in planta transformation system involved
imbibing Arabidopsis seeds overnight in an Agrobacterium culture, followed by germination (Feldmann
& Marks 1987). A large number of transgenic plants
containing T-DNA insertions were recovered, but in
general this technique has a low reproducibility.
Bechtold et al. (1993) has described a more reliable
method, in which the bacteria are vacuuminfiltrated into Arabidopsis flowers. An even simpler
technique called floral dip has become widely used
(Clough & Bent 1998). This involves simply dipping
Arabidopsis flowers into a bacterial suspension at
the time of fertilization. In both these methods, the
transformed plants are chimeric, but give rise to a
small number of transgenic progeny (typically
about 10 per plant). Similar approaches using direct
DNA transfer have been tried in other species,
but germ-line transformation has not been reproducible. For example, naked DNA has been injected
into the floral tillers of rye plants (De La Pena et al.
1987) and post-fertilization cotton flowers (Zhou
et al. 1983), resulting in the recovery of some transgenic plants. Transgenic tobacco has been produced
following particle bombardment of pollen (Touraev
et al. 1997).
An alternative to the direct transformation of germline tissue is the introduction of DNA into meristems
in planta, followed by the growth of transgenic
shoots. In Arabidopsis, this has been achieved simply
by severing apical shoots at their bases and inoculating the cut tissue with Agrobacterium suspension
(Chang et al. 1994). Using this procedure, transgenic plants were recovered from the transformed
shoots at a frequency of about 5%. In rice, explanted
meristem tissue has been transformed using Agrobacterium and particle bombardment, resulting in the
proliferation of shoots that can be regenerated into
transgenic plants (Park et al. 1996). Such procedures require only a limited amount of tissue culture.
Chloroplast transformation
So far, we have exclusively considered DNA transfer
to the plant’s nuclear genome. However, the chloroplast is also a useful target for genetic manipulation,
because thousands of chloroplasts may be present in
photosynthetic cells and this can result in levels of
transgene expression up to 50 times higher than
possible using nuclear transformation. Furthermore,
transgenes integrated into chloroplast DNA do not
appear to undergo silencing or suffer from position
effects that can influence the expression levels of
transgenes in the nuclear DNA (see Boxes 11.1 and
13.2). Chloroplast transformation also provides a
natural containment method for transgenic plants,
since the transgene cannot be transmitted through
pollen (reviewed by Maliga 1993).
The first reports of chloroplast transformation were
serendipitous, and the integration events were found
to be unstable. For example, an early experiment
in which tobacco protoplasts were cocultivated
with Agrobacterium resulted in the recovery of one
transgenic plant line, in which the transgene was
transmitted maternally. Southern-blot analysis of
chloroplast DNA showed directly that the foreign DNA
had become integrated into the chloroplast genome
(De Block et al. 1985). However, Agrobacterium does
not appear to be an optimal system for chloroplast
transformation, probably because the T-DNA complex is targeted to the nucleus. Therefore, direct
DNA transfer has been explored as an alternative
strategy. Stable chloroplast transformation was
first achieved in the alga Chlamydomonas reinhardtii,
which has a single large chloroplast occupying most
of the volume of the cell (Boynton et al. 1988).
Particle bombardment was used in this experiment.
The principles established using this simple organism were extended to tobacco, allowing the recovery
of stable transplastomic tobacco plants (Svab et al.
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Gene transfer to plants
1990b). These principles included the use of vectors
containing chloroplast homology regions, allowing
targeted integration into the chloroplast genome,
and the use of the selectable marker gene aad (aminoglycoside adenyltransferase), which confers resistance
to streptomycin and spectinomycin (Zoubenko et al.
1994). Efficient chloroplast transformation has
been achieved both through particle bombardment
(e.g. Staub & Maliga 1992) and polyethylene glycol
(PEG)-mediated transformation (Golds et al. 1993,
Koop et al. 1996). The use of a combined selectable–
screenable marker (aad linked to the gene for green
fluorescent protein) allows the tracking of transplastomic sectors of plant tissue prior to chlorophyll
synthesis, so that transformed plants can be rapidly
identified (Khan & Maliga 1999).
Plant viruses as vectors
As an alternative to stable transformation using
Agrobacterium or direct DNA transfer, plant viruses
can be employed as gene-transfer and expression
vectors. There are several advantages to the use
of viruses. First, viruses are able to adsorb to and
introduce their nucleic acid into intact plant cells.
However, for many viruses, naked DNA or RNA is
also infectious, allowing recombinant vectors to be
introduced directly into plants by methods such as
leaf rubbing. Secondly, infected cells yield large
amounts of virus, so recombinant viral vectors have
the potential for high-level transgene expression.
Thirdly, viral infections are often systemic. The virus
spreads throughout the plant, allowing transgene
expression in all cells. Fourthly, viral infections are
rapid, so large amounts of recombinant protein can
be produced in a few weeks. Finally, all known plant
viruses replicate episomally; therefore the transgenes they carry are not subject to the position
effects that often influence the expression of integrated transgenes (Box 11.1). Since plant viruses
neither integrate into nor pass through the germ
line, plants cannot be stably transformed by viral
infection and transgenic lines cannot be generated.
However, this limitation can also be advantageous
in terms of containment.
A complete copy of a viral genome can also be
introduced into isolated plant cells or whole plants
by Agrobacterium or direct DNA transfer. In this
241
manner, it is possible to generate transiently transformed cell lines or transgenic plants carrying an
integrated recombinant viral genome. In the case of
RNA viruses, transcription of an integrated cDNA
copy of the genome yields replication-competent
viral RNA, which is amplified episomally, facilitating high-level transgene expression. Transgenic
plants are persistently infected by the virus and
can produce large amounts of recombinant protein.
In the case of DNA viruses, Agrobacterium-mediated
transient or stable transformation with T-DNA
containing a partially duplicated viral genome can
lead to the ‘escape’ of intact genomes, which then
replicate episomally. The latter process, known as
‘agroinfection’ or ‘agroinoculation’, provides a very
sensitive assay for gene transfer.
As well as their use for the expression of whole
foreign proteins, certain plant viruses have recently
been developed to present short peptides on their
surfaces, similar to the phage-display technology
discussed on p. 136. Epitope-display systems based
on cowpea mosaic virus, potato virus X and tomato
bushy stunt virus have been developed as a potential
source of vaccines, particularly against animal viruses
(reviewed by Johnson et al. 1997, Lomonossoff &
Hamilton 1999).
DNA viruses as expression vectors
The vast majority of plant viruses have RNA genomes.
However, the two groups of DNA viruses that are
known to infect plants – the caulimoviruses and the
geminiviruses – were the first to be developed as vectors, because of the ease with which their small, DNA
genomes could be manipulated in plasmid vectors.
Cauliflower mosaic virus
The type member of the caulimoviruses is CaMV.
The 8 kb double-stranded DNA (dsDNA) genome
of several isolates has been completely sequenced,
revealing an unusual structure characterized by the
presence of three discontinuities in the duplex. A
map of the CaMV genome is shown in Fig. 12.15.
There are eight tightly packed genes, expressed as
two major transcripts: the 35S RNA (which essentially represents the entire genome) and the 19S
RNA (which contains the coding region for gene VI).
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CHAPTER 12
Nonessential
3‘
5‘
3‘
VII
I
IR1
19S
1
Non-essential
–
II
VI
2
8024 bp
III
35S
5‘ IR2
3 β
+
IV
α
V
VIII
As discussed earlier in the chapter, the promoter and
terminator sequences for both transcripts have been
utilized in plant expression vectors, and the 35S
promoter is particularly widely used (Box 12.1).
Only two of the genes in the CaMV genome are
non-essential for replication (gene II and gene VII)
and, since CaMV has an icosahedral capsid, the size
of the genome cannot be increased greatly without
affecting the efficiency of packaging. The maximum
capacity of the CaMV capsid has been defined as
8.3 kb and, with the removal of all non-essential
genes, this represents a maximum insert size of less
than 1 kb (Daubert et al. 1983). This restriction in
the capacity for foreign DNA represents a major limitation of CaMV vectors. Thus far it has been possible
to express a number of very small transgenes, such
as the 240 bp bacterial dhfr gene (Brisson et al.
1984), the 200 bp murine metallothionein cDNA
(Lefebvre et al. 1987) and a 500 bp human interferon cDNA (De Zoeten et al. 1989). In Chapter 10,
we describe how similar limitations were overcome
for simian virus 40 (SV40), a virus that infects
primate cells, through the development of replicon
vectors and helper viruses or complementary cell
lines supplying essential functions in trans. Unfortunately, such an approach is not possible with
Fig. 12.15 Map of the cauliflower mosaic
virus genome. The eight coding regions are
shown by the thick grey arrows, and the
different reading frames are indicated by the
radial positions of the boxes. The thin lines in
the centre indicate the (plus and minus) DNA
strands with the three discontinuities. The
major transcripts, 19S and 35S, are shown
around the outside.
CaMV, due to the high level of recombination that
occurs, leading to rapid excision of the foreign DNA
(Gronenborn et al. 1981).
Geminiviruses
Geminiviruses are characterized by their twin
(geminate) virions, comprising two partially fused
icosahedral capsids. The small single-stranded DNA
genome is circular and, in some species, is divided
into two segments called DNA A and DNA B. Interest
in geminivirus vector development was stimulated
by the discovery that such viruses use a DNA replicative intermediate, suggesting that they could be more
stable than CaMV, whose RNA-dependent replication cycle is rather error-prone (Stenger et al. 1991).
Of the three genera of geminiviruses, two have been
developed as vectors. The begomoviruses have predominantly bipartite genomes; they are transmitted
by the whitefly Bemisia tabaci and infect dicots.
Species that have been developed as vectors include
African cassava mosaic virus (ACMV) and tomato
golden mosaic virus (TGMV). The mastreviruses
have monopartite genomes; they are transmitted by
leafhoppers and predominantly infect monocots.
Species that have been developed as vectors include
POGC12 9/11/2001 11:14 AM Page 243
Gene transfer to plants
maize streak virus (MSV) and wheat dwarf virus
(WDV).
An important additional distinction between these
genera is that mastreviruses are not mechanically
transmissible. MSV, for example, has never been
introduced successfully into plants as native or
cloned DNA. Grimsley et al. (1987) were able to
overcome this problem using Agrobacterium, and
were the first to demonstrate the principle of agroinfection. They constructed a plasmid containing a
tandem dimer of the MSV genome. This dimer was
inserted into a binary vector, and maize plants were
infected with A. tumefaciens containing the recombinant T-DNA. Viral symptoms appeared within
2 weeks of inoculation. Agroinfection has been used
to introduce the genomes of a number of different
viruses into plants. It can be demonstrated that, if
the T-DNA contains partially or completely duplicated genomes, single copies of the genome can
escape and initiate infections. This may be mediated
by homologous recombination or a replicative
mechanism (Stenger et al. 1991). The study of
Grimsley et al. (1987) incidentally provided the first
evidence that Agrobacterium could transfer T-DNA
to maize. Agroinfection is a very sensitive assay for
transfer to the plant cell because of the amplification
inherent in the virus infection and the resulting
visible symptoms.
A number of geminiviruses have been developed
as expression vectors, because of the possibility of
achieving high-level recombinant-protein expression as a function of viral replication (reviewed by
Stanley 1983, Timmermans et al. 1994, Palmer &
Rybicki 1997). A generally useful strategy is the
replacement of the coat-protein gene, since this is
not required for replication and the strong promoter
can be used to drive transgene expression. In the
case of begomoviruses, which have bipartite genomes,
the coat-protein gene is located on DNA A, along
with all the functions required for DNA replication.
Replicons based on DNA A are therefore capable of
autonomous replication in protoplasts (e.g. Townsend
et al. 1986). Geminivirus replicon vectors can facilitate the high-level transient expression of foreign
genes in protoplasts. There appears to be no intrinsic
limitation to the size of the insert, although larger
transgenes tend to reduce the replicon copy number
(e.g. Laufs et al. 1990, Matzeit et al. 1991). Generally,
243
it appears that mastrevirus replicons can achieve a
much higher copy number in protoplasts than replicons based on begomiviruses. A WDV shuttle
vector capable of replicating in both E. coli and plants
was shown to achieve a copy number of greater than
3 × 104 in protoplasts derived from cultured maize
endosperm cells (Timmermans et al. 1992), whereas
the typical copy number achieved by TGMV replicons in tobacco protoplasts is less than 1000
(Kanevski et al. 1992). This may, however, reflect
differences in the respective host cells, rather than
the intrinsic efficiencies of the vectors themselves.
Geminiviruses are also valuable as expression vectors in whole plants. In the case of the mastreviruses,
all viral genes appear to be essential for systemic
infection, so coat-protein replacement vectors cannot
be used in this manner. In contrast, the coat-protein
genes of ACMV and TGMV are non-essential for systemic infection, but they are required for insect transmission (Briddon et al. 1990). Therefore, replicon
vectors based on these viruses provide an in planta
contained transient expression system. Note that
viral movement functions are supplied by DNA B, so
systemic infections occur only if DNA B is also present in the plant.
In an early study, Ward et al. (1988) replaced
most of the ACMV AV1 gene with the cat reporter
gene. In infected tobacco plants, high level CAT
activity was detected for up to 4 weeks. Interestingly, they found that deletion of the coat-protein
gene caused a loss of infectivity in plants, but this
was restored upon replacement with cat, which is
approximately the same size as the deleted gene.
This and many subsequent reports indicated that,
while there may be no intrinsic limit to the size of
replicon vectors in protoplasts, systemic infection
is dependent on preserving the size of the wildtype DNA-A component. A further limitation to
this system is that the transmissibility of the recombinant genomes is poor, probably because they
are not packaged. One way in which this can be
addressed is to generate transgenic plants in which
recombinant viral genomes are produced in every
cell. This is achieved by transforming plants with
DNA constructs containing a partially duplicated
viral genome (Meyer et al. 1992). Intact replicons can
excise from the delivered transgene in the same way
as the MSV genome escapes during agroinfection,
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CHAPTER 12
and autonomously replicating episomal copies can
be detected. Transgenic tobacco plants have also
been produced carrying an integrated copy of DNA
B (Hayes et al. 1988, 1989). In the presence of replication functions supplied by DNA A, the DNA-B
sequence is rescued from the transgene and can
replicate episomally. DNA B can then provide movement functions for DNA A, facilitating the systemic
spread of the vector.
RNA viruses as expression vectors
Most plant RNA viruses have a filamentous morphology, so the packaging constraints affecting the
use of DNA viruses, such as CaMV, should not present a limitation in vector development. However,
investigation into the use of RNA viruses as vectors
lagged behind research on DNA viruses, awaiting
the advent of robust techniques for the manipulation of RNA genomes.
A breakthrough was made in 1984, when a fulllength clone corresponding to the genome of brome
mosaic virus (BMV) was obtained. Infectious RNA
could be produced from this cDNA by in vitro
transcription (Ahlquist & Janda 1984, Ahlquist et al.
1984). The BMV genome comprises three segments:
RNA1, RNA2 and RNA3. Only RNA1 and RNA2
are necessary for replication. RNA3, which is
dicistronic, encodes the viral coat protein and movement protein. During BMV infection, a subgenomic
RNA fragment is synthesized from RNA3, containing the coat-protein gene alone. It is therefore
possible to replace the coat-protein gene with foreign DNA and still generate a productive infection
(Ahlquist et al. 1987). This was demonstrated by
French et al. (1986) in an experiment where the
coat-protein gene was substituted with the cat reporter
gene. Following the introduction of recombinant
RNA3 into barley protoplasts, along with the essential RNA1 and RNA2 segments, high-level CAT
activity was achieved.
This experiment showed that BMV was a potentially useful vector for foreign gene expression. However, to date, BMV has been used solely to study the
function of genes from other plant viruses. Following
the demonstration that infectious BMV RNA could
be produced by in vitro transcription, the genomes of
many other RNA viruses have prepared as cDNA
copies. Some of these viruses have been extensively
developed as vectors for foreign gene expression
(see comprehensive reviews by Scholthof et al. 1996,
Porta & Lomonossoff 2001). Two examples are discussed below.
Tobacco mosaic virus
TMV is one of the most extensively studied plant
viruses and was thus a natural choice for vector
development. The virus has a monopartite RNA
genome of 6.5 kb. At least four polypeptides are produced, including a movement protein and a coat
protein, which are translated from subgenomic
RNAs (Fig. 12.16). The first use of TMV as a vector
was reported by Takamatsu et al. (1987). They replaced the coat-protein gene with cat, and obtained
infected plants showing high-level CAT activity at
the site of infection. However, the recombinant
virus was unable to spread throughout the plant,
because the coat protein is required for systemic
infection.
Since there should be no packaging constraints
with TMV, Dawson et al. (1989) addressed the
deficiencies of the TMV replacement vector by generating a replication-competent addition vector, in
which the entire wild-type genome was preserved.
Dawson and colleagues added the bacterial cat gene,
controlled by a duplicated coat-protein subgenomic
promoter, between the authentic movement and
coat-protein genes of the TMV genome. In this case,
systemic infection occurred in concert with highlevel CAT activity, but recombination events in
infected plants resulted in deletion of the transgene
and the production of wild-type TMV RNA. Homologous recombination can be prevented by replacing
the TMV coat-protein gene with the equivalent
sequence from the related Odontoglossum ringspot
virus (Donson et al. 1991). This strategy has been
used to produce a range of very stable expression
vectors, which have been used to synthesize a variety of valuable proteins in plants, such as ribosomeinactivating protein (Kumagai et al. 1993) and
single-chain Fv (scFv) antibodies (McCormick et al.
1999). It has also been possible to produce complete
monoclonal antibodies by coinfecting plants with
separate TMV vectors expressing the heavy and
light immunoglobulin chains (Verch et al. 1998).
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Gene transfer to plants
245
5‘
3‘
Genomic RNA
Subgenomic
mRNA for
movement protein
Movement protein
Subgenomic
mRNA for
capsid protein
Fig. 12.16 Genome map and
expression of tobacco mosaic virus.
Capsid protein
ORF1
ORF3
AAA3’
5’cap
Genomic RNA
5835 nt
ORF2
ORF4
ORF5
Polymerase
AAA3’
5’
Subgenomic RNA
ORF5
AAA3’
5’cap
Subgenomic mRNA
(coat protein)
Fig. 12.17 Genome map and expression
of potato virus X.
Potato virus X
Potato virus X (PVX) is the type member of the
Potexvirus family. Like TMV, it has a monopartite
RNA genome of approximately 6.5 kb, which is
packaged in a filamentous particle. The genome
map of PVX is shown in Fig. 12.17, and contains
genes for replication, viral movement and the coat
protein, the latter expressed from a subgenomic
promoter. Reporter genes, such as gusA and green
fluorescent protein, have been added to the PVX
Coat protein
genome under the control of a duplicated coatprotein subgenomic promoter and can be expressed
at high levels in infected plants (Chapman et al.
1992, Baulcombe et al. 1995). As with the early
TMV vectors, there is a tendency for the transgene to
be lost by homologous recombination, but, in the
case of PVX, no alternative virus has been identified
whose coat-protein gene promoter can functionally
substitute for the endogenous viral promoter. For
this reason, PVX is generally not used for long-term
expression, but has been widely employed as a
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transient expression vector. It has been used for the
synthesis of valuable proteins, such as antibodies
(e.g. Hendy et al. 1999, Franconi et al. 1999, Ziegler
et al. 2000), and for the expression of genes that
affect plant physiology (e.g. the fungal avirulence
gene avr9) (Hammond-Kosack et al. 1995).
The stable transformation of plants with cDNA
copies of the PVX genome potentially provides a
strategy for extremely high-level transgene expression, because transcripts should be amplified to
a high copy number during the viral replication
cycle. However, instead of high-level expression,
this strategy leads to potent and consistent transgene silencing, as well as resistance to viral infection
(English et al. 1996, English & Baulcombe 1997).
The basis of this phenomenon and its implications
for transgene expression in plants are discussed
in more detail in Chapter 13. In terms of vector
development, however, it is notable that PVX-based
vectors are probably most widely used to study virusinduced gene silencing and related phenomena
(Dalmay et al. 2000) and to deliberately induce
silencing of homologous plant genes (reviewed by
Baulcombe 1999).
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CHAPTER 13
Advances in transgenic
technology
Introduction
In the previous three chapters, we discussed the
early development of transgenic technology, which
focused on the evolution of reliable techniques for
gene transfer and transgene expression in animals
and plants. The standard use of this technology was
to add genetic information to the genome, generating a gain of function in the resulting cells and transgenic organisms.
Now that such gene-transfer processes are routine, the focus of transgenic technology has shifted
to the provision of more control over the behaviour
of transgenes. New techniques have evolved in
parallel in animals and plants. Advances have
come about through the development of inducible
expression systems that facilitate external regulation of transgenes and the exploitation of site-specific
recombination systems to make precise modifications in target genomes. In mice, the combination
of gene targeting, site-specific recombination and
inducible transgene expression makes it possible to
selectively switch on and off both transgenes and
endogenous genes in a conditional manner. Other
routes to gene silencing have also been explored,
such as the expression of antisense RNA and the
recently described phenomenon of RNA interference.
Transgenic animals and plants are also being used
increasingly as tools for the analysis of genomes.
We discuss the development of tagging systems for
the rapid cloning of genes disrupted by insertional
mutagenesis and the use of entrapment vectors,
which allow the high-throughput analysis of gene
expression and function.
Inducible expression systems
In many gene-transfer experiments, the production
of a recombinant protein is the ultimate goal, in
which case it is generally appropriate to maximize
transgene expression using a strong constitutive
promoter. However, there are also situations in
which maximized transgene expression is undesirable. For example, if a recombinant protein is toxic,
constitutive high-level expression would be lethal
and would prevent the recovery of stably transformed cell lines. In other experiments, the timing of
transgene expression is critical. This would be the
case if we wished to study the effect of protein overexpression at a specific point in the cell cycle in cultured cells or at a particular developmental stage in
transgenic animals and plants. These issues can be
addressed through the use of inducible expression
systems, in which transgene expression is controlled
by an external stimulus.
Endogenous inducible promoters
A number of inducible expression systems have been
developed based on promoters from endogenous
cellular or viral genes. An early example is the
Drosophila heat-shock promoter. Most cells respond
to elevated temperature by synthesizing heat-shock
proteins, which include molecular chaperons and
other proteins with protective functions (Parsell &
Lindquist 1983). The response is controlled at the
level of transcription by a heat-labile transcription
factor, which binds to heat-responsive promoters
in the corresponding genes. The promoter of the
Drosophila hsp70 gene has been widely used to drive
transgene expression, both in Drosophila itself (Lis
et al. 1983) and in heterologous systems (e.g. Wurm
et al. 1986). In transgenic flies, any gene linked to
the hsp70 promoter is more or less inactive at room
temperature, but high-level expression in all cells
can be induced by heating to 37°C for about 30 min.
The heat-shock promoter is unusual in that the
stimulus that activates it is physical. Most inducible
promoters respond to chemicals, which must be
supplied to the transformed cells or transgenic
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CHAPTER 13
organisms in order to activate the expression of a
linked transgene. In mammals, several promoters
are known to be activated by glucocorticoid hormones or synthetic analogues, such as dexamethasone. Two of these have been extensively used for
inducible transgene expression – the mouse metallothionein promoter (Hager & Palmiter 1981) and
the long-terminal-repeat (LTR) promoter of mouse
mammary tumour virus (MMTV) (Lee et al. 1981).
The metallothionein promoter is also induced by
interferons and heavy metals, such as cadmium and
zinc, allowing the transgene to be activated in transgenic animals, e.g. by including a source of heavy
metals in the drinking-water. An example of zincinduced activation of a human growth-hormone
gene in transgenic mice is discussed on p. 208. A
metal-inducible expression system has also been
developed for plants, although the components of
the system are derived from yeast (Mett et al. 1993).
Endogenous inducible promoters used in plants
include the PR-1a promoter, which is responsive to
benzothiadiazole (Gorlach et al. 1996).
There are several limitations associated with the
use of endogenous inducible promoters, and most
systems based on such promoters suffer from one or
more of the following problems. First, they tend to be
somewhat leaky, i.e. there is a low to moderate level
of background transcription in the absence of induction. Secondly, the level of stimulation achieved
by induction (the induction ratio) is often quite low,
typically less than 10-fold. Thirdly, there are often
cytotoxic side-effects, caused by the activation of
other endogenous genes that respond to the same
inductive stimulus. The inducers of many of the
commonly used endogenous promoters are also
toxic if contact is prolonged. Fourthly, the kinetics of
induction are often not ideal. For example, in transgenic animals and plants, there may be differential
uptake of the inducer into different cell types and
it may be eliminated slowly. There has therefore
been great interest in the development of alternative
inducible expression systems.
Recombinant inducible systems
Many of the disadvantages of endogenous inducible
promoters can be addressed using recombinant
systems. In the ideal system, induction is dependent
on the supply of a non-toxic agent that specifically
activates the transgene without interacting with
any endogenous promoters. The agent is taken up
rapidly and evenly, but has a short half-life, allowing
rapid switching between induced and non-induced
states. A range of inducers might be available with
different properties. Steps towards this ideal system
have been achieved using promoters and transcription factors that are either heterologous in the
expression host or completely artificial.
The lac and tet repressor systems
The first heterologous systems were based on bacterial control circuits (Fig. 13.1). Hu and Davidson
(1987) developed an inducible expression system
for mammalian cells, incorporating the essential
elements of the lac repressor control circuit. In
Escherichia coli, transcription of the lac operon is
switched off in the absence of lactose by a repressor
protein encoded by the gene lacI. This protein binds
to operator sites, the most important of which lies just
downstream from the promoter, and thus inhibits
transcriptional initiation. In the presence of lactose
or a suitable analogue, such as isopropyl-β-dthiogalactoside (IPTG), the lac repressor undergoes a
conformational change that causes it to be released
from the operator sites, allowing transcription to
commence.
In order to use the lac circuit in eukaryotic cells,
Hu and Davison (1987) modified the lacI gene by
adding a eukaryotic initiation codon, and then made
a hybrid construct in which this gene was driven by
the Rous sarcoma virus (RSV) LTR promoter. The
construct was introduced into mouse fibroblasts and
a cell line was selected that constitutively expressed
the lac repressor protein. This cell line was transiently transfected with a number of plasmids containing the cat reporter gene, driven by a modified
simian virus 40 (SV40) promoter. Each of these plasmids also contained a lac operator site somewhere
within the expression construct. The investigators
found that, when the operator sites were placed in
the promoter region, transcription from the reporter
construct was blocked. However, transcription could
be derepressed by supplying the cells with IPTG,
resulting in strong chloramphenicol transacetylase
(CAT) activity.
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Advances in transgenic technology
249
Repressor gene
P
No induction
With induction
I
R
R
P
Target gene
P
O
Component
Lac system
Tet system
Repressor gene
lac I
tet R
A similar system, based on the E. coli tetracycline
operon, was developed for tobacco plants (Gatz &
Quail 1988). The tet operon is carried on a bacterial
transposon that confers resistance to the antibiotic
tetracycline. Similarly to the lac system, the tet operon
is switched off by a repressor protein, encoded by the
tetR gene, which binds to operator sites around the
promoter and blocks transcriptional initiation.
Tetracycline itself binds to this repressor protein and
causes the conformational change that releases
the tet operon from repression. Since tetracycline
inhibits bacteria at very low concentrations, the tet
repressor has a very high binding constant for the
antibiotic, allowing derepression in the presence of
just a few molecules. The tet repressor also has
very high affinity for its operator sites. Therefore, cell
cultures and transgenic tobacco plants expressing
TetR were able to inhibit reporter gene expression
from a cauliflower mosaic virus (CaMV) 35S promoter surrounded by three tet operator sites. This
repressed state could be lifted rapidly by the application of tetracycline (Gatz et al. 1991).
The lac and tet repressor systems both show
minimal background transcription in the presence
of the appropriate repressor protein, and a high
induction ratio is therefore possible. In the lac system
the maximum induction ratio is approximately 50,
whereas in the tet system up to 500-fold induction
has been achieved (Figge et al. 1988, Gatz et al.
I
Target gene
O
Fig. 13.1 Summary of repressionbased inducible expression-control
circuits based in the lac and tet operons
of E. coli.
R
R
Lac repressor
Tet repressor
I
IPTG
tetracycline/doxycycline
O
lac operator
tet operator
1992). Remarkably, the bacterial repressor proteins
appear quite capable of interacting with the eukaryotic transcriptional apparatus and functioning as
they do in bacteria, despite the many mechanistic
differences in transcriptional control between prokaryotes and eukaryotes.
The tet activator and reverse activator systems
A disadvantage of repressor-based systems is that, in
order to function effectively, high-level constitutive
expression of repressor molecules is required to
suppress background transgene activity. However,
both the LacI and TetR proteins are cytotoxic at high
levels.
To address these problems, TetR and LacI have
been converted into activators by generating fusion
proteins, in which the repressor is joined to the herpes
simplex virus (HSV) VP16 transactivator (Labow
et al. 1990, Gossen & Bujard 1992). In these systems,
only the DNA-binding specificity of the repressor
proteins is exploited. The binding of the modified
bacterial proteins to operator sites within the transgene leads to transcriptional activation, because the
VP16 protein acts positively on the transcriptional
apparatus. The operator sites have effectively become
enhancers and the inducers (IPTG and tetracycline)
have effectively become repressors (Fig. 13.2). The
tet transactivator (tTA) system has been more
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CHAPTER 13
tTA
P
–Tet
+Tet
TET
t TA
t TA
t TA
P
Target gene
Target gene
tetO
P
tetO
Fig. 13.2 The tet transactivator system
(tTA).
rt TA
P
–Tet
rt TA
Target gene
P
tetO
rt TA
+Tet
TET
rt TA
P
Target gene
tetO
Fig. 13.3 The reverse tet transactivator
system (rtTA).
widely used than the equivalent lac system, because
very high levels of IPTG are required to inhibit LacI
binding in mammalian cells and this is toxic (Figge
et al. 1988). Many different proteins have been produced in mammalian cell lines using the tTA system,
particularly cytotoxic proteins, whose constitutive
expression would rapidly lead to cell death (Wu &
Chiang 1996). In cells, a low background activity
has been reported and an induction ratio of approximately 105 can be achieved. However, toxic effects
of prolonged tetracycline exposure have been reported
in transgenic animals, as well as unequal uptake
of tetracycline into different organs, resulting in
fluctuating basal transcription levels and cellspecific effects (reviewed by Saez et al. 1997).
A further modification to the tet system has led
to marked improvements. A mutated form of the tTA
protein has been generated whose DNA-binding
activity is dependent on rather than abolished by
tetracycline (Gossen et al. 1995). This protein is
called reverse tTA (rtTA) and becomes an activator
in the presence of tetracycline. In this system, the
antibiotic is once again an inducer, but there is no
requirement for prolonged exposure (Fig. 13.3). An
early example of the use of this system is described by
Bohl et al. (1997). Myoblasts were transformed with
the rtTA system using a retroviral vector in which
the erythropoietin complementary DNA (cDNA) was
placed under the control of a tetracycline-inducible
promoter. These cells were implanted into mice,
and erythropoietin secretion could be controlled by
feeding the mice doxycycline, a derivative of tetracycline with a shorter half-life. An important finding
was that long-term control of the secretion of this
hormone was possible, with important implications
for the use of inducible expression systems for gene
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Advances in transgenic technology
therapy. A number of convenient plasmid- and virusbased vectors have been developed, allowing the
components of the rtTA system to be transferred to
the expression host in one experiment (for example,
see Hofmann et al. 1996, Paulus et al. 1996, Schultze
et al. 1996). A reverse-transactivator lac system has
also been developed, with an induction ratio of
approximately 104 (Baim et al. 1991).
Heterologous use of steroids: Drosophila
ecdysone in mammals and mammalian
glucocorticoids in plants
Steroid hormones are lipophilic molecules that penetrate cells rapidly and are eliminated within a few
hours. The use of heterologous steroids for inducible
transgene expression is therefore advantageous,
because, in addition to their favourable kinetics,
such molecules should not activate endogenous
signalling pathways in the expression host and
should therefore have limited toxicity.
Ecdysone is a steroid hormone found only in
insects and is responsible for the extensive morphological changes that occur during moulting. As with
other steroid-like signalling molecules, the hormone
acts through a heterodimeric transcription factor of
the nuclear receptor family. In Drosophila, this receptor comprises the products of the genes ecdysone
receptor (ecr) and ultraspiracle (usp). The hormone
and its signalling pathway are not found in mammalian cells. Therefore, transgenes including an
ecdysone response element in the promoter can be
induced by exogenously supplied ecdysone or its
analogue, muristerone A, in cells or transgenic animals expressing the components of the Drosophila
receptor. The unmodified Drosophila system has a
poor induction ratio, but this can be improved using
chimeric receptors and mammalian components
(Yao et al. 1992, 1993). In a significant improvement, No and colleagues were able to achieve an
induction ratio of 104 by generating a hybrid system
in which the ecdysone receptor gene was expressed
as a fusion with the HSV VP16 transactivator, and the
ultraspiracle protein was replaced with a mammalian
homologue, the retinoid X receptor. Background
activity was reduced to near zero by altering the
DNA sequence recognized by the hybrid receptor
(No et al. 1996).
251
Although glucocorticoid induction has been used
as an endogenous inducible expression system in
mammalian cells, mammalian glucocorticoid receptors do not function properly in transgenic plants.
However, the ligand-binding domain can be expressed as a fusion with other transcription factors
and used to bring the activity of such transcription
factors under inducible control. A potentially very
powerful system has been described in which the
glucocorticoid-receptor steroid-binding domain has
been fused to the DNA-binding domain of the yeast
transcription factor GAL4 and the VP16 transactivation domain. In this system, a CaMV 35S promoter
modified to contain six GAL4-recognition sites is
used to drive transgene expression. Genes placed
under the control of this promoter can be induced
100-fold in the presence of dexamethasone (Aoyama
& Chua 1997). An ecdysone-based system for plants
has also been described, in which an agrochemical
acts as the inducer, such that chemicals used in
the field could function as inducers (Martinez et al.
1999).
Chemically induced dimerization
A novel strategy for inducible transgene regulation
has been developed, exploiting essentially the same
principles as the yeast two-hybrid system (p. 169).
This technique, termed chemically induced dimerization (CID), involves the use of a synthetic divalent
ligand to simultaneously bind and hence bring
together separate DNA-binding and transactivation
domains to generate a functional transcription factor. The initial system utilized the immunosuppressant drug FK-506. This binds with high specificity
to an immunophilin protein called FKBP12, forming
a complex that suppresses the immune system by
inhibiting the maturation of T cells (reviewed by
Schreiber 1991). For transgene induction, an artificial
homodimer of FK-506 was created, which could
simultaneously bind to two immunophilin domains.
Therefore, by expressing fusion proteins in which
the GAL4 DNA-binding domain and the VP16
transactivator were each joined to an immunophilin
domain, the synthetic homodimer could recruit a
functional hybrid transcription factor capable of
activating any transgene carrying GAL4 recognition elements (Belshaw et al. 1996).
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CHAPTER 13
FK– 506
Specific interactions
I
Cyclosporin A
Immunophilin
Synthetic dimer
C
Cyclophilin
FK–506/Cyclosporin A
Chimeric constructs
Immunophilin GAL4
Cyclophilin
GAL4
Immunophilin
P
P
GAL4
VP16
I
– CID
I
GAL4
P
VP16
Cyclophilin
+ CID
I
Target gene
C
C
GAL4 VP16
P
Since this homodimer can also recruit nonproductive combinations (e.g. two GAL4 fusions),
an improved system has been developed using an
artificial heterodimer specific for two different
immunophilins (Belshaw et al. 1996). In this case,
FK-506 was linked to cyclosporin A, a drug that
binds specifically to a distinct target, cyclophilin.
This heterodimer was shown to effectively assemble
a transcription factor comprising an FKBP12–GAL4
fusion and a cyclophilin–VP16 fusion, resulting
in strong and specific activation of a reporter gene
in mammalian cells (Fig. 13.4). A more versatile
system has been developed that exploits the ability
of another immunosupressant drug, rapamycin, to
mediate the heterodimerization of FKBP12 and a
kinase known as FRAP (Rivera et al. 1996). In this
system, FKBP12 and FRAP are each expressed as
fusions with the components of a functional transcription factor. In the absence of rapamycin there
is no interaction between these fusions, but when
the drug is supplied they assemble into a hybrid
Target gene
Fig. 13.4 Overview of chemically
induced dimerization between the
synthetic FK-506/cyclosporin A
conjugate and fusion proteins
containing immunophilin and
cyclosporin domains. Dimerization
assembles a functional transcription
factor that can activate a promoter with
GAL4 response elements.
transcription factor that can activate transgene
expression. Transgenic mice containing a growthhormone gene controlled by a CID-regulated promoter showed no expression in the absence of the
inducer, but high levels of human growth hormone
24 h after induction with rapamycin (Magari et al.
1997). The advantage of this system is that rapamycin
is rapidly taken up by cells in vivo, and it decays
rapidly. The major disadvantage of immunosuppressant drugs as chemical inducers of dimerization
is their pharmacological side-effects. An active area
of current research is the design of modified CID
systems that do not interact with endogenous
targets in the immune system, and thus provide
rapid and effective transgene induction with no
side-effects (Liberles et al. 1997).
Inducible protein activity
The inducible expression systems discussed above
are all regulated at the level of transcription, such
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Advances in transgenic technology
253
Oestrogen
Tamoxifen
Hsp90
OR
Hsp90
OR
Fig. 13.5 Principle of the oestrogeninducible expression system. The
oestrogen receptor (OR) is normally
sequestered by heat-shock protein 90
(Hsp90). However, in the presence of
oestrogen or its analogue Tamoxifen,
the receptor is released and can form
dimers and activate target genes.
that there is often a significant delay between induction and response and between removal of induction
and return to the basal state. Where a rapid response
to induction is critical, inducible systems that operate
at the post-translational level can be utilized. For
example, the mammalian oestrogen receptor exists
in an inert state in the absence of oestrogen, because
the hormone-binding domain interacts with heatshock protein 90 (Hsp90) to form an inactive complex.
When oestrogen is present, it binds to its receptor,
causing a conformational change that releases the
receptor from Hsp90. The receptor is then free to
dimerize and interact with DNA (Fig. 13.5). In principle, any protein expressed as a fusion with the
oestrogen-binding domain will similarly interact
with Hsp90 and form an inactive complex (Picard
1994). A recombinant protein can thus be expressed
at high levels in an inactive state, but can be
activated by feeding cells or transgenic animals
with oestrogen or an analogue, such as Tamoxifen,
which does not induce endogenous oestrogenresponsive genes (Littlewood et al. 1995). A similar
system has been devised using a mutant-form
progesterone receptor, which can no longer bind
progesterone but can be induced with the antiprogestin RU486 (Garcia et al. 1992, Vegeto et al.
1992). An induction ratio of up to 3500 has been
demonstrated in transgenic mice and, importantly,
the inductive response occurs when the drug is
supplied at a dose more than 100-fold below that
required for it to function as an antiprogestin (Wang
et al. 1997a,b).
OR OR
Target gene
OR OR
Target gene
Applications of site-specific
recombination
Until recently, there was no generally applicable
method for the precise in vivo manipulation of DNA
sequences in animal and plant genomes. In mice,
gene targeting by homologous recombination allows
specific changes to be introduced into preselected
genes, but it had proved impossible to extend the
technique to other animals or to plants. Furthermore,
even gene targeting is limited by the fact that the
targeted gene is modified in the germ line; thus all cells
in the mouse are similarly affected from the beginning
of development and throughout its entire lifetime.
Over the last 10 years, general methods have
become available that allow in vivo transgene manipulation in any animal or plant species. Importantly,
by using such methods in concert with inducible or
cell-type-specific expression systems, it is possible to
generate transgenic organisms in which transgenes
can be conditionally modified. In mice, the use of
these methods in combination with gene targeting
allows the production of conditional mutants (conditional knockouts), in which an endogenous gene is
inactivated specifically in certain cell types or at a
particular stage of development. These methods are
based on a specialized genetic process, termed sitespecific recombination.
Site-specific recombination
Site-specific recombination differs from homologous
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Box 13.1 Visible marker genes
Reporter genes are widely used for in vitro assays of
promoter activity (Box 10.1). However, reporters that
can be used as cytological or histological markers are
more versatile, because they allow gene expression
profiles to be determined in intact cells and whole
organisms.
b-galactosidase and b-glucuronidase
The E. coli lacZ gene encodes b-galactosidase, an
enzyme that hydrolyses b-d-galactopyranosides, such
as lactose, as well as various synthetic analogues. Like
CAT, b-galactosidase activity can be assayed in vitro,
although with the advantage that the assays are
non-radioactive. For example, cell lysates can be
assayed spectrophotometrically using the chromogenic
substrate ONPG*, which yields a soluble yellow
compound (Norton & Coffin 1985). Alternatively, a
more sensitive fluorometric assay may be preferred,
using the substrate MUG*. For histological staining,
the substrate X-gal* yields an insoluble blue
precipitate that marks cells brightly. The lacZ gene
was first expressed in mammalian cells by Hall et al.
(1983) to confirm transfection. For these experiments,
the gene was linked to the SV40 early promoter and
the mouse mammary tumour virus (MMTV) LTR
promoter. Fusions between the hsp70 promoter
and lacZ were also constructed and shown to drive
heat-shock-inducible b-galactosidase expression in
Drosophila (Lis et al. 1983). One disadvantage of
lacZ as a marker is that certain mammalian cells,
and many plants, show a high level of endogenous
b-galactosidase activity, which can obscure the
analysis of chimeric genes (Helmer et al. 1984).
The E. coli gusA gene, which encodes the enzyme
b-glucuronidase (GUS), is an acceptable alternative
(Jefferson et al. 1986). This marker is preferred in
plants, due to the minimal background activity of the
endogenous enzyme (Jefferson et al. 1987a), but has
also been used successfully in animals (e.g. Jefferson
et al. 1987b). Similar in vitro and histological assay
formats to those described for b-galactosidase are
also available for GUS, e.g. a histochemical substrate,
X-gluc*, which yields an insoluble blue precipitate.
Luciferase
CAT, GUS and b-galactosidase are stable proteins,
which persist in the cells that express them. One
problem with stable reporter proteins is that, while
they provide useful markers for gene activation, they
are less useful for assaying transcriptional repression
or rapid changes in gene activity. Luciferase was
introduced as a novel reporter gene in 1986, for
use in both plants (Ow et al. 1986) and animals
(De Wet et al. 1987). The original marker gene,
luc, was isolated from the North American firefly
Photinus pyralis and encoded a single polypeptide of
550 amino acids. The enzyme catalyses the oxidation
of luciferin, in a reaction requiring oxygen, ATP
and the presence of magnesium ions. When excess
substrate is supplied, a flash of light is emitted that is
proportional to the amount of enzyme present. This
can be detected using a luminometer, a scintillation
counter as a luminometer or even photographic film
(Wood & DeLuca 1987). Important advantages of
the luciferase system include its very high sensitivity
(more than 100-fold more sensitive than lacZ) and
the rapid decay of light emission. Luciferase has
therefore been used to analyse the activity of genes
with oscillating expression profiles, such as the
Drosophila period gene (Brandes et al. 1996). The
amenability of the luciferase system has been
expanded by the isolation of alternative luciferases
from other organisms, which bioluminesce in different
colours (e.g. see Thompson et al. 1990). A bacterial
luciferase gene, luxA, has also been used as a marker
in transgenic plants (Koncz et al. 1987).
Green fluorescent protein
The most recent addition to the growing family of
reporters is green fluorescent protein (GFP), from the
jellyfish Aequoria victoria. Over the last 5 years, this
remarkable molecule has emerged as one of the most
versatile tools in molecular and cellular biology and
is being used to investigate an increasing variety of
biological processes in bacteria, yeast, animals and
plants (reviewed by Tsien 1998, Haseloff et al.
continued
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Advances in transgenic technology
255
Box 13.1 continued
1999, Ikawa et al. 1999, Naylor 1999). GFP is a
bioluminescent marker that causes cells to emit
bright green fluorescence when exposed to blue or
ultraviolet light. However, unlike luciferase, GFP has
no substrate requirements and can therefore be used
as a vital marker to assay cellular processes in real
time. Other advantages of the molecule include the
fact that it is non-toxic, it does not interfere with
normal cellular activity and it is stable even under
harsh conditions (Ward & Bokman 1982).
GFP was first used as a heterologous marker
in Caenorhabditis elegans (Chalfie et al. 1994).
However, early experiments with GFP expression in
a variety of other organisms, including Drosophila
(Wang & Hazelrigg 1994), mammalian cell lines
(Marshall et al. 1995) and plants (Haseloff &
Amos 1995, Hu & Chen 1995, Sheen et al. 1995),
identified a number of difficulties in the heterologous
expression of the gfp gene. Modifications have been
necessary for robust GFP expression in some plants
(Chiu et al. 1996). In Arabidopsis, for example, the
original gfp gene is expressed very poorly due to
aberrant splicing. This problem was addressed by
removing a cryptic splice site recognized in this plant
(Haseloff et al. 1997). The original gfp gene has
been extensively modified to alter various properties
of the protein, such as the excitation and emission
wavelengths, to increase the signal strength and to
reduce photobleaching (e.g. Heim & Tsein 1996,
recombination in several important respects. In
terms of gene manipulation, the most important
differences between these processes concern the
availability of the recombinase and the size and
specificity of its target sequence. Homologous recombination is a ubiquitous process that relies on
endogenous recombinase enzymes present in every
cell, whereas site-specific recombination systems are
very specialized and different systems are found
in different organisms. Homologous recombination
occurs between DNA sequences with long regions of
homology but no particular sequence specificity,
whereas site-specific recombination occurs at short,
specific recognition sites. This means that target
Zolotukhin et al. 1996, Cormack et al. 1997). As a
result, a range of variant GFPs are available which
can be used for dual labelling (e.g. Tsien & Miyawaki
1998; reviewed by Ellenberg et al. 1999). Fluorescent
proteins of other colours are also available, including
a red fluorescent protein from Anthozoa (Matz et al.
1999). A mutant form of this protein changes from
green to red fluorescence over time, allowing it to
be used to characterize temporal gene expression
patterns (Terskikh et al. 2000).
GFP is particularly useful for generating fusion
proteins, providing a tag to localize recombinant
proteins in the cell. This facilitates the investigation
of intracellular protein trafficking, and even the
transport of proteins between cells. An early example
of this application was the use of GFP to monitor the
movement of ribonucleprotein particles during
oogenesis in Drosophila (Wang & Hazelrigg 1994).
Kohler et al. (1997) have used GFP to study the
exchange of molecules between plant organelles,
while Wacker et al. (1997) have investigated the
transport of a GFP-tagged protein along the secretory
pathway. The use of GFP to study the real-time
dynamics of a systemic viral infection in plants
was described by Padgett et al. (1996).
* Abbreviations: ONPG O-nitrophenyl-β-d-galactopyranoside;
MUG 4-methylumbelliferyl-β-d-galactoside; X-gal:
5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside; X-gluc:
5-bromo-4-chloro-3-indolyl-β-d-glucuronic acid.
sites for site-specific recombination can be introduced easily and unobtrusively into transgenes, but
recombination will only occur in a heterologous cell
if a source of recombinase is also supplied. The power
of site-specific recombination as a tool for genome
manipulation thus relies on the ability of the experimenter to supply the recombinase enzyme on a conditional basis.
A number of different site-specific recombination
systems have been identified and several have been
studied in detail (reviewed by Craig 1988, Sadowski
1993). Some recombinases, such as bacteriophage
λ integrase, require various accessory proteins for
efficient recombination. However, the simplest systems
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require only the recombinase and its target sequence.
Of these, the most extensively used are Cre recombinase from bacteriophage P1 (Lewanodski & Martin
1997) and FLP recombinase (flippase) from the
2 µm plasmid of the yeast Saccharomyces cerevisiae
(Buchholz et al. 1998). These have been shown to
function in many heterologous eukaryotic systems
including mammalian cells and transgenic animals
and plants (reviewed by Sauer 1994, Ow 1996,
Metzger & Feil 1999). Both recombinases recognize
34 bp sites (termed loxP and FRP, respectively) comprising a pair of 13 bp inverted repeats surrounding
an 8 bp central element. FRP possesses an additional
copy of the 13 bp repeat sequence, although this has
been shown to be non-essential for recombination.
Cre recombinase has been used most extensively in
mammals, because it works optimally at 37°C. The
optimal temperature for FLP recombinase is 30°C
(Buchholz et al. 1996).
Site-specific deletion of transgene
sequences
Site-specific deletion of unwanted transgenes
The reaction catalysed by Cre recombinase is shown
in Fig. 13.6. If two loxP sites are arranged as direct
repeats, the recombinase will delete any intervening
DNA, leaving a single loxP site remaining in the
genome. If the loxP sites are arranged as inverted
repeats, the intervening DNA segment is inverted.
Both reactions are reversible. However, when the
loxP sites are arranged in the same orientation,
excision is favoured over reintegration, because the
excised DNA fragment is rapidly degraded.
The ability of flanking loxP sites to delineate any
sequence of interest for site-specific deletion has
numerous applications. The most obvious of these is
the deletion of unwanted sequences, such as marker
genes. This approach has been used, for example, as
a simplified strategy to generate targeted mutant
mice containing point mutations. Recall from Chapter 11 that traditional strategies for generating subtle
mutants in mice involve two rounds of homologous
recombination in embryonic stem cells (ES cells)
(p. 207). Such strategies are very inefficient, because
homologous recombination is a rare event. However,
in the Cre recombinase-based approach, a second
round of homologous recombination is unnecessary
(Kilby et al. 1993). As shown in Fig. 13.7, gene targeting is used to replace the wild-type allele of a
given endogenous gene with an allele containing a
point mutation, and simultaneously to introduce
markers, such as neo and Tk, for positive and negative
selection. The positive–negative selection markers
within the homology region are flanked by loxP sites.
A T A A CT T CGT A T A A T GT A T GCT A T A CGA A GT T A T
T A T T GA A GCA T A T T A CA T A CGA T A T GCT T CA A T A
CRE
+
CRE
Fig. 13.6 Structure of the loxP site and reactions
catalysed by Cre recombinase when paired loxP
sites, shown as arrows, are arranged in different
orientations.
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Fig. 13.7 Gene targeting followed by
marker excision, catalysed by Cre
recombinase. Initially, positive and
negative markers (neo and Tk), flanked
by loxP sites, are introduced by
homologous recombination (a second
negative marker, in this case encoding
diphtheria toxin, is included outside the
homology region to eliminate random
integration events). Following selection
for neo on G418, Cre recombinase is
used to excise both markers, leaving
a single loxP site remaining in the
genome. The excision event can be
identified by selection for the absence
of Tk, using ganciclovir or FIAU.
Asterisk indicates mutation.
1
2
3
>
2
257
4
5
>
NEO
TK
3*
Locus
Targeting
construct
DIPA
Homologous recombination
select on G418
>
1
>
NEO
2
TK
3*
4
5
Targeted
locus
CRE-mediated recombination
select on ganciclovir/FIAU
>
1
A second negative marker (e.g. the gene for diphtheria
toxin) is included outside the homology region to
select against random integration events. Cells that
have lost the diphtheria-toxin gene and survive selection for neo are likely to represent authentic targeting events. Such cells are then transfected with a
plasmid expressing Cre recombinase, which catalyses the excision of the remaining markers, leaving
a clean point mutation and no evidence of tinkering
except for a single loxP site remaining in one intron.
Negative selection using ganciclovir or 1-(2-deoxy2-fluoro-β-d-arabinofuranosyl)-5-iodouracil (FIAU)
identifies cells that have lost the markers by selection
against Tk.
Similar strategies can be used to remove marker
genes from plants, as first demonstrated by Dale and
Ow (1991). These investigators used Agrobacterium
to transform tobacco-leaf explants with a CaMV 35Sluciferase reporter construct. The transfer DNA
(T-DNA) also contained a selectable marker for
hygromycin resistance, flanked by loxP sites. Transgenic plants were regenerated under hygromycin
selection and leaf explants from these plants were then
transformed with a second construct, in which Cre
recombinase was driven by the CaMV 35S promoter.
This construct also contained the nptII gene and
the second-round transgenic plants were selected
on kanamycin. Ten of the 11 plants tested were
found to be hygromycin-sensitive, even though they
continued to express luciferase, showing that the
original marker had been excised. Since the cre/nptII
construct was introduced separately, it was not
2
4
3*
5
Clean
replacement
linked to the original T-DNA and segregated in
future generations, leaving ‘clean’ transgenic plants
containing the luciferase transgene alone.
Site-specific transgene activation and switching
While commonly used as a method to inactivate
transgenes by deletion, site-specific recombination
can also activate transgenes or switch between
the expression of two transgenes (Fig. 13.8). In one
method, termed recombinase-activated gene expression
(RAGE), a blocking sequence, such as a polyadenylation site, is placed between the transgene and
its promoter, such that the transgene cannot be
expressed. If this blocking sequence is flanked by
loxP sites, Cre recombinase can be used to excise the
sequence and activate the transgene.
P
poly A
Target gene
+ Cre recombinase
P
Target gene
Fig. 13.8 Overview of the recombinase activation of gene
expression (RAGE) strategy. A polyadenylation signal is
inserted between the promoter and target gene, blocking its
expression. However, if this signal is flanked by loxP sites, Cre
recombinase can be used to excise the block, bringing the
promoter and gene into juxtaposition and thus activating
gene expression.
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This strategy was first used in transgenic mice to
study the effect of SV40 T-antigen expression in
development (Pichel et al. 1993). In this case, Cre
recombinase was expressed under the control of a
developmentally regulated promoter. Essentially the
same strategy was used in transgenic tobacco plants
to activate a reporter gene in seeds (Odell et al.
1994). In this case, Cre recombinase was expressed
under the control of a seed-specific promoter. An
important feature of both these experiments was the
use of two separate transgenic lines, one expressing
Cre recombinase in a regulated manner and one
containing the target gene. Crosses between these
lines brought both transgenes together in the hybrid
progeny, resulting in the conditional activation of
the transgene based on the expression profile of
Cre. This is an extremely versatile and widely used
strategy, because it allows ‘mix and match’ between
different Cre transgenic and ‘responder’ lines. We
return to this subject below.
Site-specific transgene integration
Site-specific integration of transgenes can occur if
the genome contains a recombinase recognition
site. This may be introduced by random integration
or (in mice) by gene targeting. Using an unmodified
Cre–loxP system, transgene integration occurs at a
low efficiency, because, as discussed above, the equilibrium of the reaction is shifted in favour of excision.
Initial attempts to overcome this problem by providing transient Cre activity had limited success (see
Sauer & Henderson 1990, Baubonis & Saur 1993).
However, high-efficiency Cre-mediated integration
has been achieved in plants (Albert et al. 1995) and
mammalian cells (Feng et al. 1999) using mutated
or inverted loxP sites. Site-specific transgene integration into mammalian cells has also been achieved
using FLP recombinase (O’Gorman et al. 1991).
Transgene integration by site-specific recombination has many advantages over the random
integration that is normally achieved by illegitimate
recombination. For example, if a region of the genome
can be identified that is not subject to negative
position effects (Box 11.1), transgenic lines with a
loxP site at this position can be used for the stable
and high-level expression of any transgene (e.g.
Fukushige & Sauer 1992). Also, due to the precise
nature of site-specific recombination, transgenic loci
generated by this method are likely to be less complex than loci generated by random integration.
Chromosome engineering
Site-specific recombination between widely separated
target sites or target sites on different chromosomes
can be used to generate large deletions, translocations
and other types of chromosome mutation. Chromosome engineering by site-specific recombination
was first reported by Golic (1991), using FLP recombinase in Drosophila, but similar experiments have
now been carried out in plants and mice. Precise intrachromosomal deletions can be generated in mice by
two rounds of gene targeting, introducing loxP sites
at distant sites, followed by Cre-mediated recombination (Ramirez-Solis et al. 1995, Li et al. 1996). In
plants, where gene targeting is very inefficient, an
ingenious scheme has been developed where loxP
sites are arranged in tandem on a transformation
construct, one inside a Ds transposon and one outside (Fig. 13.9). The transposon is placed between a
marker gene and its promoter. When this construct is
introduced into tobacco plants containing the autonomous transposon Ac to provide a source of transposase, the Ds element can excise from the transgene,
as revealed by marker-gene expression. In most
heterologous plants, Ac–Ds elements reintegrate at a
position that is linked to the original site. Although
the site of reintegration cannot be controlled, this
nevertheless defines a large chromosomal segment
that can be excised by Cre recombinase (Medberry
et al. 1995, Osbourne et al. 1995). Translocations
are more difficult to engineer, because interchromosomal site-specific recombination is inefficient, and
inventive selection strategies are required to identify
the desired products (e.g. see Qin et al. 1994, Smith
et al. 1995, Van Deursen et al. 1995).
Cre-mediated conditional
mutants in mice
In mice, gene targeting and site-specific recombination can be used in a powerful combined approach to
generate conditional knockout mutants. Essentially,
targeting vectors are designed so that part of a
selected endogenous gene becomes flanked by loxP
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Advances in transgenic technology
P
Fig. 13.9 Chromosome engineering
in plants with Cre recombinase. A
construct integrates into the plant
genome. The construct contains a
promoter separated from the reporter
gene gusA by a Ds transposon
containing a loxP site. Another loxP site
is located just upstream of the promoter.
The addition of transposase (e.g. by an
Ac element elsewhere in the genome)
causes the Ds element to excise, thereby
bringing the gusA gene adjacent to the
promoter. The excision event can thus
be identified by the onset of GUS activity
in the plant. The Ds element tends to
insert a few tens of kilobases away
on the same chromosome. Cre
recombinase can then delete the large
genomic region between the two
loxP sites.
sites, or floxed. The usual strategy is to insert the loxP
sites into introns flanking an essential exon, since
this generally does not interfere with the normal
expression of the gene. Cre recombinase is then
supplied under the control of a cell-type-specific,
developmentally regulated or inducible promoter,
causing the gene segment defined by the loxP sites
to be deleted in cells or at the developmental stage
specified by the experimenter. This addresses a
major limitation of traditional gene-knockout techniques, i.e. that, if the mutation has an embryonic
lethal phenotype, only its earliest effects can be
investigated.
The general methodology for such experiments,
as we discussed earlier, is to cross two lines of
transgenic mice, one carrying the floxed target gene
and the other carrying the conditional cre transgene. As the number of reports of such experiments
has increased, more and more transgenic mouse
lines are becoming available, with Cre expressed
under the control of different conditional promoters.
For example, a mouse line with Cre expressed specifically in the lens of the eye was generated by Lasko
et al. 1992. Lines are also available with Cre expressed
specifically in the mammary gland (Wagner et al.
1997) and developing sperm (O’Gorman et al. 1997).
Lines in which Cre is expressed in germ cells or in
early development are known as ‘deleter’ lines and
259
DS
gus A
Transformation
construct
Integration
P
DS
P
gus A
gus A
GUS activty
+ transposase
DS
+ Cre recombinase
are used to remove marker genes and generate Cremediated constitutive gene knockouts.
In the first examples of the conditional knockout
approach, Gu et al. (1994) generated a Cre transgenic line expressing the recombinase under the
control of the lck promoter, such that it was
expressed only in T cells. This strain was crossed to
targeted mice in which part of the DNA polymerase
β gene was floxed, leading to T-cell-specific excision
of an essential exon. Kuhn et al. (1995) mutated
the same gene, but they used the metallothionein
promoter to express Cre recombinase, allowing induction of site-specific recombination with interferon.
Although successful, this experiment highlighted
many of the inadequacies of inducible promoters.
There was pronounced variation in the efficiency
of excision in different tissues, probably reflecting
differential uptake of the inducer. Furthermore,
high-level background activity of Cre was observed
in the spleen, resulting in excision of the gene segment in the absence of induction, probably caused
by the presence of endogenous interferons. The tTA
system has been used to bring Cre expression under
the control of tetracycline administration, although
a high level of background activity was also observed
in this experiment, resulting in excision of the target
gene prior to induction (St Ogne et al. 1996). Tighter
control has been possible using post-translational
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induction. For example, Cre has been expressed as a
fusion with the ligand-binding domain of the oestrogen receptor (Fiel et al. 1996). When this transgene
was crossed into an appropriate responder strain,
the background excision was minimal and Cre was
strongly induced by Tamoxifen. Several strains of
Cre mice are now available, in which Tamoxifenor RU486-induced site-specific recombination has
been shown to be highly efficient (e.g. Brocard et al.
1997, Wang et al. 1997a, Schwenk et al. 1998).
Further transgenic strategies
for gene inhibition
Traditional gene-transfer strategies add new genetic
information to the genome, resulting in a gain of
function phenotype conferred by the transgene. Gene
targeting and site-specific recombination now provide
us with the ability to disrupt or delete specific parts
of the mouse genome, allowing loss-of-function
phenotypes to be studied, but this approach cannot be used in other animals or in plants. A range
of alternative, more widely applicable transgenic
strategies have therefore been developed for gene
inhibition. These strategies involve the introduction
of new genetic information into the genome, but,
instead of conferring a gain of function, the transgene interferes with the expression of an endogenous gene, at either the RNA or the protein level. The
actual target gene is not affected. The resulting loss
of function effects are termed functional knockouts or
phenocopies.
Gene inhibition at the RNA level
Antisense RNA transgenes
Antisense RNA has the opposite sense to mRNA.
The presence of complementary sense and antisense
RNA molecules in the same cell can lead to the
formation of a stable duplex, which may interfere
with gene expression at the level of transcription,
RNA processing or possibly translation (Green et al.
1986). Antisense RNA is used as a natural mechanism
to regulate gene expression in a number of prokaryote
systems (Simons & Kleckner 1988) and, to a lesser
extent, in eukaryotes (e.g. Kimelman & Kirchner
1989, Lee et al. 1993, Savage & Fallon 1995).
Transient inhibition of particular genes can be
achieved by directly introducing antisense RNA or
antisense oligonucleotides into cells. However, the
transformation of cells with antisense transgenes
(in which the transgene is inverted with respect to
the promoter) allows the stable production of antisense RNA and thus the long-term inhibition of gene
expression. This principle was established in transgenic animals and plants at about the same time.
Katsuki et al. (1988) constructed an expression
cassette in which the mouse myelin basic protein
(MBP) cDNA was inverted with respect to the
promoter, thus producing antisense RNA directed
against the endogenous gene. In some of the transgenic mice, there was up to an 80% reduction in the
levels of MBP, resulting in the absence of myelin
from many axons and generating a phenocopy of
the myelin-depleted ‘shiverer’ mutation. Smith et al.
(1988) generated transgenic tomato plants carrying an antisense construct targeting the endogenous polygalacturonase (pg) gene. The product of this
gene is an enzyme that causes softening and leads to
overripening. The levels of pg mRNA in transgenic
plants were reduced to 6% of the normal levels and
the fruit had a longer shelf-life and showed resistance to bruising.
Antisense constructs have been widely used in
transgenic animals and plants for gene inhibition.
However, the efficiency of the technique varies widely
and the effects can, in some cases, be non-specific.
In some experiments, it has been possible to shut
down endogenous gene activity almost completely,
as demonstrated by Erickson et al. (1993), who
used an inverted cDNA to generate antisense RNA
against the mouse wnt-1 gene and reduced endogenous mRNA levels to 2% of normal. Conversely,
Munir et al. (1990) designed a construct to generate
antisense RNA corresponding to the first exon and
intron of the mouse Hprt gene, and observed no
reduction in endogenous mRNA levels at all, even
though the presence of antisense RNA was confirmed. The level of inhibition apparently does not
depend on the size of the antisense RNA or the part of
the endogenous gene to which it is complementary.
For example, Moxham et al. (1993) achieved a 95%
reduction in the level of Gαi2 protein through the
expression of antisense RNA corresponding to only
39 bp of the gene’s 5′ untranslated region.
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Conditional gene silencing can be achieved by
placing antisense constructs under the control of
an inducible promoter. The expression of antisense
c-myc under the control of the MMTV LTR promoter
resulted in the normal growth of transformed cells
in the absence of induction, but almost complete
growth inhibition in the presence of dexamethasone
(Sklar et al. 1991). Experiments in which the tTA system was used to control antisense expression in plants
have also been reported (e.g. Kumar et al. 1995).
Ribozyme constructs
Ribozymes are catalytic RNA molecules that carry
out site-specific cleavage and (in some cases) ligation reactions on RNA substrates. The incorporation
of ribozyme catalytic centres into antisense RNA
allows the ribozyme to be targeted to particular
mRNA molecules, which are then cleaved and
degraded (reviewed by Rossi 1995, James & Gibson
1998). An important potential advantage of ribozymes
over antisense RNA is that ribozymes are recycled
after the cleavage reaction and can therefore inactivate many mRNA molecules. Conversely, antisense
inhibition relies on stoichiometric binding between
sense and antisense RNA molecules.
The use of ribozyme constructs for specific gene
inhibition in higher eukaryotes was established in
Drosophila. In the first such report, Heinrich et al.
(1983) injected Drosophila eggs with a P-element
vector containing a ribozyme construct targeted
against the white gene. They recovered transgenic
flies with reduced eye pigmentation, indicating that
expression of the endogenous gene had been inhibited. A ribozyme construct has also been expressed
under the control of a heat-shock promoter in
Drosophila (Zhao & Pick 1983). In this case, the
target was the developmental regulatory gene fushi
tarazu ( ftz). It was possible to generate a series of
conditional mutants with ftz expression abolished at
particular stages of development, simply by increasing the temperature to 37°C.
Ribozymes have also been used in mammalian
cell lines, predominantly for the study of oncogenes
and in attempts to confer resistance to viruses
(reviewed by Welch et al. 1998). There has been
intensive research into ribozyme-mediated inhibition
of HIV, and remarkable success has been achieved
261
using retroviral vectors, particularly vectors carrying multiple ribozymes (reviewed by Welch et al.
1998, Muotri et al. 1999). So far, there have been
relatively few reports of ribozyme expression in
transgenic mice. Larsson et al. (1994) produced mice
expressing three different ribozymes targeted against
β2-macroglobulin mRNA, and succeeded in reducing endogenous RNA levels by 90%. Tissue-specific
expression of ribozymes has also been reported. A
ribozyme targeted against glucokinase mRNA was
expressed in transgenic mice under the control of
the insulin promoter, resulting in specific inhibition
of the endogenous gene in the pancreas (Efrat et al.
1994). Recently, retroviral delivery of anti-neuregulin
ribozyme constructs into chicken embryos has been
reported (Zhao & Lemke 1998). Inhibition of neuregulin expression resulted in embryonic lethality,
generating a very close phenocopy of the equivalent
homozygous null mutation in mice.
Cosuppression
Cosuppression refers to the ability of a sense transgene to suppress the expression of a homologous
endogenous gene. This surprising phenomenon was
first demonstrated in plants, in a series of experiments designed to increase the levels of an endogenous protein by introducing extra copies of the
corresponding gene. In an attempt to increase the
amount of pigment synthesized by petunia flowers,
Napoli et al. (1990) produced transgenic petunia
plants carrying multiple copies of the chalcone
synthase (chs) gene. This encodes an enzyme that
converts coumaroyl-CoA and 3-malonyl-CoA into
chalcone, a precursor of anthocyanin pigments. The
presence of multiple transgene copies was expected
to increase the level of enzyme and result in deeper
pigmentation. However, in about 50% of the plants
recovered from the experiment, exactly the opposite
effect was observed, i.e. the flowers were either pure
white or variegated with purple and white sectors.
Similar findings were reported by Van der Krol et al.
(1988) using a transgene encoding another pigmentbiosynthesis enzyme, dihydroflavonol-4-reductase.
In both cases, it appeared that integration of multiple copies of the transgene led to the suppression of
some or all of the transgenes and the cosuppression
of homologous endogenous genes.
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While troublesome in terms of generating plant
lines with high transgene expression levels, cosuppression can also be exploited as a tool for specific
gene inactivation. There have been many reports of
this nature. For example, transgenic tomatoes have
been produced containing a partial copy of the pg
gene in the sense orientation (Smith et al. 1990). As
with the antisense pg transgenic plants generated
previously by the same group (see above), strong
inhibition of the endogenous gene was achieved,
resulting in fruit with a prolonged shelf-life. Cosuppression has also been demonstrated in animals
(Pal-Bhadra et al. 1997, Bahramian & Zabl 1999,
Dernberg et al. 2000) and is probably related to a
similar phenomenon called quelling, which has been
described in fungi (reviewed by Selker 1997, 1999).
The mechanism of cosuppression in plants is
complex and can involve silencing at either the
transcriptional or post-transcriptional levels (for
details, see Box 13.2). One of the most remarkable
Box 13.2 Transgene silencing, cosuppression and RNA interference
Transgene silencing is a complex phenomenon,
occurring in all eukaryotes, caused by the
introduction of foreign nucleic acid into the cell.
Typically, expression of the affected transgene is
reduced or abolished, associated with increased
methylation at the transgenic locus. An
understanding of transgene silencing is important,
first, because it is a serious impediment to the use
of animal and plant systems for the expression of
foreign genes and, second, because if a transgene
is homologous to an endogenous gene, the
endogenous gene can also be silenced by
cosuppression. Several different forms of silencing
can be distinguished.
Position-dependent silencing and
sequence-dependent silencing
These forms of silencing can affect single-copy
transgenes and are not, therefore, homologydependent. Position-dependent silencing occurs
where a transgene integrates into a genomic region
containing heterochomatin. The repressive chromatin
structure and DNA methylation can spread into the
transgenic locus from the flanking genomic DNA
(Matzke & Matzke 1998); therefore silencing results
from a negative position effect (position effects are
discussed in more detail in Box 11.1). Single-copy
transgenes may also be silenced, even if they
integrate into a genomic region that lacks negative
position effects. For example, integrated retrovirus
vectors often undergo de novo silencing associated
with increased levels of DNA methylation (Jahner
et al. 1982) and, indeed, this methylation may
spread outwards into flanking host DNA and
inactivate linked genes (Jahner & Jaenisch 1985).
Many unrelated transgenes in animals and plants
have been subject to this type of silencing,
suggesting that a specific sequence is not
responsible. It is possible that eukaryotic genomes
possess mechanisms for scanning and identifying
foreign DNA sequences, perhaps based on their
unusual sequence context, and then inactivating
them by methylation (Kumpatla et al. 1998).
Prokaryotic DNA may be recognized in this manner,
since prokaryotic sequences act as a strong trigger
for de novo methylation, e.g. in transgenic mice
(Clark et al. 1997).
Homology-dependent gene silencing (HDGS)
HDGS is caused by the presence of multiple
transgene copies (reviewed by Gallie 1998, Grant
1999, Plasterk & Ketting 2000, Hammond et al.
2001). The suppression of transgene expression can
occur at the transcriptional or post-transcriptional
levels, and homologous endogenous genes are often
cosuppressed. Single-copy transgenes homologous
to an endogenous gene may, in some cases, also be
sufficient to induce cosuppression. In transcriptional
gene silencing (TGS), no mRNA is produced from the
silenced genes and affected loci act as nucleation
points for heterochromatin formation and DNA
methylation. In post-transcriptional gene silencing
(PTGS), transcription is actually required for silencing
to take place, and it induces the degradation of
mRNA, so that very little accumulates in the
continued
POGC13 9/11/2001 11:13 AM Page 263
Box 13.2 continued
loxP
loxP
loxP
loxP
Cre
loxP
Fig. B13.1 An experiment to demonstrate homology-dependent gene silencing in mammals (also called repeat-induced
gene silencing (RIGS)). A transgene construct containing the human b-globin cDNA was modified to contain a single loxP
site, which is recognized by Cre recombinase. Transgenic mice were generated carrying multiple copies of the transgene,
and in these animals the locus was highly methylated and b-globin expression was low. However, when Cre recombinase
was expressed, recombination between the loxP sites resulted in the excision of all copies of the transgene except one.
Reduction in the transgene copy number resulted in increased expression, accompanied by reduced methylation at the
transgenic locus. (After Garrick et al. 1998.)
cytoplasm. The severity of silencing often correlates
with transgene copy number, and this can be
demonstrated directly through the use of site-specific
recombination, as shown in the figure above.
Different processes may trigger homologydependent silencing, including the ability of
homologous DNA sequences to form secondary
structures and the synthesis of aberrant RNA
molecules, leading to the production of dsRNA. Both
processes are stimulated if the transgenic locus is
complex, particularly if transgenes or partial
transgenes are arranged as inverted repeats. There
may be some cross-talk between the TGS and PTGS
mechanisms, but this is currently unclear. In plants,
transgenes carried by RNA viruses can induce PTGS,
a phenomenon that has been termed virus-induced
gene silencing (VIGS). Only replication-competent
viruses have this effect, providing further evidence
that dsRNA is the trigger for this process. The highlevel expression of sense transgenes can also induce
PTGS, and it has been suggested that the plant cell
may recognize such aberrant transcripts and convert
them into dsRNA.
PTGS and RNAi as a common mechanism
– the evidence
The involvement of dsRNA in many cases of PTGS in
plants suggests that the underlying basis of this
phenomenon may be similar to RNA interference
(RNAi), which is also a post-transcriptional process.
As discussed in the text, both PTGS and RNAi are
systemic, i.e. silencing can spread from the site at
which it was induced throughout the entire organism.
Recently, small RNA molecules, called guide RNAs,
have been identified in plants affected by PTGS and
animals subjected to RNAi, which can also spread
systemically (Hamilton & Baulcombe 1999,
Hammond et al. 2000, Parrish et al. 2000). It is
proposed that these RNAs may assemble with certain
proteins to generate a catalytic endonuclease
complex that cleaves target mRNA molecules
efficiently and in a homology-dependent fashion. In
cases of PTGS, guide RNAs may form from aberrant
transcripts generated by integrated transgenes or
directly from dsRNA viral genomes.
What is silencing for?
It is possible that both homology-dependent and
sequence-dependent silencing have evolved as a
defence against ‘invasive’ nucleic acids (Yoder et al.
1997, Jones et al. 1998, Jensen et al. 1999, Li et al.
1999). This has been supported by the recent
isolation of mutants in several organisms that show
deficiencies in PTGS or RNAi. Animals impaired
for RNAi show increased rates of transposon
mobilization, whereas plants impaired for PTGS
are more susceptible to viral infection. Interestingly,
similar gene products have been identified in diverse
organisms, providing further evidence for a link
between PTGS and RNAi. A comprehensive discussion
of this exciting new area of research is outside the
scope of this book, but the interested reader can
consult several excellent reviews on the subject
(Plasterk & Ketting 2000, Hammond et al. 2001).
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aspects of post-transcriptional gene silencing (PTGS)
is that it is a systemic phenomenon, suggesting that
a diffusible signal is involved. This can be demonstrated by grafting a non-silenced transgenic scion
on to a silenced transgenic host. The silencing effect
is able to spread into the graft, and the systemic effect
works even if the two transgenic tissues are separated by up to 30 cm of wild-type stem (Palauqui
et al. 1997, Voinnet et al. 1998).
PTGS in plants can be induced not only by
integrated transgenes but also by RNA viruses, as
long as there is a region of homology between the
virus genome and an integrated gene. For example,
the virus may carry a sequence that is homologous
to an endogenous gene or to a transgene integrated
into the host genome. The effect also works if the
plant is transformed with a cDNA construct corresponding to part of the virus genome, as demonstrated by Angell and Baulcombe (1997). The
rationale behind this experiment was to transform
plants with a cDNA construct corresponding to a
chimeric potato virus X (PVX) genome containing
the gusA reporter gene. Expression of the transgene was expected to generate very high levels of
β-glucuronidase (GUS) activity, because, after transcription of the transgene, the resulting viral RNA
would be amplified by the virus’s own replication
system. However, disappointingly, all of the transgenic plants produced extremely low levels of viral
RNA and GUS activity. The plants also showed an
absence of PVX symptoms and were resistant to PVX
infection. The virus-induced silencing effect only
worked using replication-competent vectors, suggesting that the double-stranded RNA (dsRNA) intermediate involved in viral replication was the trigger
for silencing (see Box 13.2).
Such is the efficiency with which PVX RNA can
silence homologous genes in the plant genome that
PVX vectors have been used very successfully to
generate functional knockouts in plants (reviewed
by Baulcombe 1999). For example, Burton et al.
(2000) described the infection of tobacco plants with
PVX vectors containing a cDNA sequence putatively
encoding a cellulose synthase. The inoculated plants
showed a dwarf phenotype, and levels of cellulose in
affected leaves were reduced by 25%. On the basis of
this evidence, the investigators concluded that the
cDNA did indeed encode such an enzyme and was
capable of cosuppressing the endogenous cellulose
synthase gene.
RNA interference
RNA interference (RNAi) is a novel phenomenon
that has the potential to become an extremely
powerful tool for gene silencing in any organism.
The process was discovered by Fire et al. (1998)
while investigating the use of antisense and sense
RNA for gene inhibition in the nematode worm
Caenorhabditis elegans. In one experiment, they introduced both sense and antisense RNA into worms
simultaneously and observed a striking and specific
inhibitory effect, which was approximately 10-fold
more efficient than either single RNA strand alone.
Further investigation showed that only a few
molecules of dsRNA were necessary for RNAi in C.
elegans, indicating that, like the action of ribozymes,
the effect was catalytic rather than stoichiometric.
Interference could be achieved only if the dsRNA
was homologous to the exons of a target gene, indicating that it was a post-transcriptional process. The
phenomenon of RNAi appears to be quite general,
and this strategy has been used more recently for
gene silencing in many other organisms, including Drosophila (Kennerdell & Carthew 2000), mice
(Wianny & Zernicka-Goetz 2000) and plants (Waterhouse et al. 1998, Chuang & Meyerowitz 2000). In
C. elegans, RNAi is now the standard procedure for
gene inactivation, and it is becoming increasingly
favoured in Drosophila and plants, due to its potency
and specificity. Direct injection of the RNA is unnecessary. The use of a construct containing adjacent
sense and antisense transgenes producing hairpin
RNA (e.g. Chuang & Meyerowitz 2000, Tavernarakis
et al. 2000) or a single transgene with dual opposing
promoters (Wang et al. 2000) provides a stable
source of dsRNA and hence the potential for permanent gene inactivation. Like PTGS in plants,
RNAi is also systemic. RNAi-mediated silencing
can be achieved in C. elegans by injecting dsRNA into
any part of the body (Fire et al. 1998), but, more
remarkably, simply placing worms in a solution
containing dsRNA or feeding them on bacteria
that synthesize dsRNA is sufficient to trigger the
effect (Tabara et al. 1998, Timmons & Fire 1998).
The similarities between RNAi and PTGS in plants
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Advances in transgenic technology
suggest a common molecular basis, which we discuss briefly in Box 13.2.
Gene inhibition at the protein level
Intracellular antibodies
Antibodies bind with great specificity to particular
target antigens and have therefore been exploited in
many different ways as selective biochemical agents.
Examples discussed in this book include the immunological screening of cDNA expression libraries
(p. 109) and the isolation of recombinant proteins
by immunoaffinity chromatography (p. 76). The
microinjection of antibodies into cells has been
widely used to block the activity of specific proteins,
but the limitation of this approach is that the
inhibitory effect is transient (reviewed by Morgan
& Roth 1988). Specific inhibitory effects can also
be achieved by microinjecting cells with RNA from
hybridoma cell lines (Valle et al. 1982, Burke &
Warren 1984). Such experiments provided the first
evidence that non-lymphoid cells can synthesize
and assemble functional antibodies.
To achieve long-term inhibition of specific proteins,
cells can be transformed with cDNA constructs that
allow the expression of intracellular antibodies (sometimes termed intrabodies) (Richardson & Marasco
1995). An important consideration here is that
antibodies are large multimeric proteins with, in
addition to antigen binding, various effector functions that are non-essential for intracellular protein
inhibition. The strategy for expressing intracellular antibodies has been radically simplified using
modified antibody forms, such as single-chain Fv
(scFv) fragments (Fig. 13.10). These comprise the
antigen-binding variable domains of the immunoglobulin heavy and light chains, linked by a flexible
peptide arm. Such fragments retain the specificity of
Heavy chain constant region
Heavy chain variable region
Light chain variable region
265
the parent monoclonal antibody, but are encoded by
a single, relatively small transgene. Further modifications to the expression construct allow the antibody
to be targeted to particular intracellular compartments, such as the nucleus, mitochondria or cytosol.
It should be noted, however, that antibodies are
normally folded and assembled in the endoplasmic
reticulum (ER) and Golgi apparatus and are generally less stable in cell compartments outside the
secretory pathway.
Due to their long half-life in the ER, intracellular
antibodies have been particularly useful for the
inhibition of cell-surface receptors, which pass through
this compartment en route to the plasma membrane.
For example, the cell-surface presentation of functional interleukin-2 (IL2) receptors was completely
abolished in Jurkat cells stably expressing an antiIL2Rα scFv fragment in the ER, rendering these cells
insensitive to exogenously applied IL2 (Richardson
et al. 1995). More recently, the same result has
been achieved using lentivirus vectors expressing
the scFv fragment, demonstrating how intracellular antibodies can be valuable for gene therapy
(Richardson et al. 1998). Intracellular antibodies
have also been used to abolish the activity of oncogenes (Beerli et al. 1994, Cochet et al. 1998, Caron
de Fromentel et al. 1999) and to confer virus resistance by inhibiting replication (reviewed by Rondon
& Marasco 1997). Functional antibodies, both fullsized immunoglobulins and fragments, can also be
expressed in plants. Hiatt et al. (1989) were the first
to demonstrate the expression of plant recombinant
antibodies, dubbed ‘plantibodies’, and subsequent
experiments have shown that this strategy can be
used, as in animal cells, to combat viral diseases by
targeting specific viral proteins (Conrad & Fielder
1998). Antibodies expressed in plants have also
been used to interfere with physiological processes
in the plant, e.g. antibodies against abscisic acid have
been used to disrupt signalling by this hormone in
tobacco (Artsaenko et al. 1995). Note that plants
can also be used as bioreactors to produce recombinant antibodies for therapeutic use (Chapter 14).
Disulphide bond
Flexible peptide linker
IgG
scFv
Fig. 13.10 Comparison of a normal immunoglobulin
molecule with a single-chain Fv fragment.
Dominant-negative mutants
In diploid organisms, most loss-of-function mutations
generate recessive or semidominant (dosage-related)
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phenotypes, because the remaining wild-type copy
of the gene provides enough gene product for normal or near-normal activity. However, some loss-offunction mutations are fully dominant over the
wild-type allele, because the mutant gene product
interferes with the activity of the wild-type protein.
Such mutants are known as dominant negatives, and
principally affect proteins that form dimers or larger
multimeric complexes.
The deliberate overexpression of dominantnegative transgenes can be used to swamp a cell
with mutant forms of a particular protein, causing
all functional molecules to be mopped up into
inactive complexes. The microinjection of DNA constructs or in vitro-synthesized dominant-negative
RNA into Xenopus embryos has been widely used to
examine the functions of cell-surface receptors in
development, since many of these are dimeric (e.g.
see Amaya et al. 1991, Hemmati-Brivanlou &
Melton 1992). Dominant-negative proteins stably
expressed in mammalian cells have been used predominantly to study the control of cell growth and
proliferation. A dominant-negative ethylene receptor
from Arabidopsis has been shown to confer ethylene
insensitivity in transgenic tomato and petunia. The
effects of transgene expression included delayed fruit
ripening and flower senescence (Wilkinson et al.
1997).
Transgenic technology for
functional genomics
In the last 5 years, the complete sequences of the
genomes of several important model eukaryotic
species have been published: the yeast S. cerevisiae
(Mewes et al. 1997), the nematode worm C. elegans
(C. elegans Sequencing Consortium 1998), the fruit
fly Drosophila melanogaster (Adams et al. 2000), the
model plant Arabidopsis thaliana (Arabidopsis Genome
Initiative 2000) and, most recently, the human
genome itself (International Human Genome
Sequencing Consortium 2001, Venter et al. 2001).
With the wealth of information that has been generated by these genome projects, the next important
step is to find out what all the newly discovered
genes actually do. This is the burgeoning field of
functional genomics, which aims to determine the
function of all the transcribed sequences in the
genome (the transcriptome) and all the proteins that
are made (the proteome).
In other parts of the book, we have already discussed a number of functional genomics techniques,
including microarrays (p. 116) and variants of the
yeast two-hybrid system (p. 169). However, perhaps
the most important way to establish a gene’s function is to see what happens when that gene is either
mutated or inappropriately expressed in the context
of the whole organism. This chapter has focused
on the development of novel transgenic strategies
for the analysis of gene function in animals and
plants, but such approaches are only applicable to
individual genes. For functional genomics, technologies must be available that allow the highthroughput analysis of gene function, which is the
only way we can begin to understand what the
10 000–40 000 genes in the genomes of higher
eukaryotic cells are for. Such technologies are
discussed below.
Insertional mutagenesis
Traditional techniques for generating mutations
involve the use of radiation or chemical mutagens.
These tend to generate point mutations, and isolating the genes corresponding to a particular mutant
phenotype can be a laborious task, particularly in
the large genomes of vertebrates and plants.
An alternative is to use DNA as a mutagen. The
genomes of most animals and plants contain transposable elements – DNA elements that have the ability
to jump from site to site in the genome, occasionally
interrupting genes and causing mutations. Some of
the gene-silencing methods discussed in the previous section appear to be based on defence strategies,
many involving DNA methylation, that exist to resist
the movement of such elements in the genome. However, if these endogenous transposable elements can
be mobilized at a sufficient frequency, they can be
used to deliberately interrupt functional genes and
generate insertional mutants. Importantly, populations of animals or plants carrying transposable
elements can be set up for saturation mutagenesis,
such that a suitable number of transposition events
is induced to theoretically interrupt each gene in the
genome at least once somewhere in the population
(e.g. see Walbot 2000).
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Endogenous transposable elements are not the
only sequences that cause insertional mutagenesis.
Foreign DNA introduced into a cell may occasionally integrate into an existing gene and disrupt its
expression. Insertional mutants have therefore
been identified as by-products of other gene-transfer
experiments. As an example, we consider the study
of Yokoyama et al. (1993). They introduced a transgene encoding the enzyme tyrosinase into the male
pronuclei of fertilized mouse eggs and recovered a
number of founders that went on to produce transgenic lines. In one line, they found a unique and
unexpected phenotype – situs inversus (the reversal
of left–right asymmetry among the developing organs)
and inversion of embryonic turning. This phenotype
had nothing to do with tyrosinase activity, since the
other lines were not similarly affected, but represented
integration of the transgene into a locus required for
correct specification of the left–right axis. Generally,
aberrant results such as these are not reported
unless they warrant investigation in their own right,
so insertional mutants are likely to be produced in
many unrelated transgenic experiments. A number
of serendipitous cases of insertional mutagenesis
in transgenic mice, caused by either pronuclear
microinjection or retroviral transfer of foreign DNA,
have been reviewed (Rijkers et al. 1994).
Gene tagging
The use of transposable elements or transgenes
for insertional mutagenesis is advantageous over
traditional mutagenesis methods, because the
interrupted locus becomes ‘tagged’ with a unique
sequence. Cloning genes tagged with endogenous
transposons can sometimes be complicated if there
are many copies of the element in the genome, but
the use of heterologous transposons or transgenes
can provide a truly unique sequence tag that identifies the interrupted gene. A genomic library can
therefore be generated from the mutant, and the interrupted gene can be identified using hybridization or
the polymerase chain reaction (PCR) to identify clones
containing the tag. Such clones will also contain
flanking DNA from the interrupted genomic locus.
Compared with chromosome walking and similar
techniques (p. 107), tagging is a simple procedure
that homes in directly on the mutant gene. Several
267
Digest DNA with restriction
endonuclease, to cut at sites
marked
Circularize, ligate
Primer 1
Primer 2
Outwardly-facing primers
anneal at core region,
PCR amplification
Fig. 13.11 Inverse PCR. The core region is indicated by
the wavy line. Restriction sites are marked with arrows,
and the left and right regions which flank the core region
are represented by closed and open boxes. Primers are
designed to hybridize with core sequences and are extended
in the directions shown. PCR amplification generates a linear
fragment containing left and right flanking sequences.
PCR-based techniques can be used to directly amplify
genomic DNA flanking a transposon or transgene
tag, thus avoiding the necessity for library construction in phage-λ or cosmid vectors (reviewed by Maes
et al. 1999). The inverse PCR strategy is shown in
Fig. 13.11 (Ochman et al. 1988). Genomic DNA from
the mutant is digested with a restriction enzyme for
which there is no site in the insertional tag. The DNA
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is thus divided into fragments, one of which will
contain the entire insertion sequence flanked by
genomic DNA tails. The tails, however, have compatible sticky ends and in a suitably dilute ligation
reaction can be self-ligated to generate a circular
genomic fragment. PCR primers annealing at the
edges of the transposon tag but facing outwards can
then be used to amplify the flanking genomic
sequences. Such PCR strategies have been used to
generate fragment libraries of flanking sequences
for the rapid identification of genes associated with
particular phenotypes (Parinov et al. 1999, Tissier
et al. 1999). If appropriate, these products can be
used to screen traditional genomic or cDNA libraries
to isolate the uninterrupted clone.
An even greater simplification of this procedure is
possible if the origin of replication from a bacterial
plasmid and an antibiotic-resistance gene are
included in the tag (Perucho et al. 1980a). In this
case, self-ligation of the construct as above generates an autonomously replicating plasmid containing genomic DNA. If whole preparations of genomic
DNA are used to transform competent bacterial cells,
only the tag-derived plasmid will replicate, while
other genomic circles will be rapidly diluted from the
culture. Furthermore, only bacteria carrying the
antibiotic-resistance marker will survive selection,
allowing pure plasmid DNA corresponding to the
flanking regions of the genomic insert to be automatically prepared. This plasmid-rescue technique is
illustrated in Fig. 13.12.
Both endogenous transposable elements and
transgene insertions have therefore been used to
isolate genes by tagging in a variety of species. In
D. melanogaster, P elements have been widely used
because of the high transposition frequency and the
availability of strains that lack P elements, allowing
lines containing individual insertions of interest to
be derived by back-crossing (Bingham et al. 1981).
Tc1 elements have been used for transposon tagging
in C. elegans (e.g. Moerman et al. 1986), Ac–Ds
elements have been widely used in maize (reviewed
by Gierl & Saedler 1992) and Tam elements have
been used in the snapdragon (Antirrhinum majus).
Tam elements are particularly useful because the
transposition frequency can be increased more than
800-fold if the temperature at which the plants are
raised is reduced to 15°C (Coen et al. 1989). Also, it
ori Ampr
Digest with restriction
endonuclease, cut out
sites marked
ori Ampr
Circularize, ligate
ori Ampr
Transform E. coli
selection
Colonies contain rescued plasmid of interest
Fig. 13.12 The principle of plasmid rescue, a technique for
isolating genomic DNA sequences flanking a transposon or
transgene integration site.
is notable that approximately 5% of all naturally
occurring recessive mutations in the mouse are
caused by a particular family of retroviruses. The
cloning of several mouse genes has been facilitated
by virtue of their linkage to a proviral sequence
(e.g. Bowes et al. 1993).
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Advances in transgenic technology
The transposition of Drosophila P elements appears
to be restricted to drosophiloid insects (Handler et al.
1993), but other transposable elements are more
promiscuous. The Ac–Ds elements of maize have
been shown to function in a wide range of plants –
dicots as well as monocots – and have been extensively used as insertional mutagens and gene tags
in heterologous species (reviewed by Sundaresan
1996). T-DNA can also be used as a tag following
transfer by Agrobacterium tumefaciens. T-DNA mutagenesis and tagging has been used in Arabidopsis
(Feldmann 1991, Krysan et al. 1999) and has recently
been extended to other plant systems, such as rice
(Jeon et al. 2000).
Vectors for insertional mutagenesis and tagging
The development of vectors specifically for insertional
mutagenesis and gene tagging arose directly from
the use of transposons and the recovery of serendipitous insertional mutants in gene-transfer experiments. The transposable elements of several species
have been developed as vectors. Modifications have
been carried out in order to control the number of
insertion events and to facilitate the cloning and
analysis of tagged genes. The use of recombinant Pelement derivatives for controlled transposon mutagenesis in Drosophila is a good example (reviewed by
Cooley et al. 1988). Two fly strains are involved. One
strain contains the mutator, a defective P element
carrying useful marker genes, which can be mobilized when provided with a source of transposase.
A second strain contains the jumpstarter, a wingsclipped element, which provides transposase in trans
but lacks the cis-acting elements required for its
own mobilization. During a controlled mutagenesis
screen, a single jumpstarter element is crossed into a
mutator-containing strain, whereupon transposition
occurs. In subsequent generations the mutator is
stabilized when the chromosome carrying the mutator
element segregates from the chromosome bearing
the jumpstarter element. The inclusion of a marker
gene in the mutator P element allows screening or
selection for transformed flies and facilitates subsequent manipulation of the modified locus. The inclusion of a bacterial selectable-marker gene and an
E. coli origin of replication allows recovery of the
tagged gene by plasmid rescue, as discussed above.
269
Entrapment constructs
Entrapment constructs are genetic tools that combine three important principles of gene-transfer
technology: (i) the random integration of transgenes
causes insertional mutagenesis and tags the mutated
gene; (ii) randomly integrated DNA sequences are
subject to variable position effects (Box 11.1); and
(iii) reporter genes can be used to assay the activity
of regulatory elements to which they become joined
(Box 10.1). Entrapment constructs are insertional
mutagenesis vectors adapted to provide information
about the genomic region into which they integrate.
Such vectors contain a reporter gene, whose expression is activated by regulatory elements in the
surrounding DNA. When they integrate into the
genome, the pattern of reporter-gene expression
reveals the activity of nearby genes, allowing investigators to screen for appropriate or interesting
insertion events. The corresponding gene can be
cloned, because it is tagged by the insertion.
Enhancer traps in Drosophila
The prototype entrapment construct was the Drosophila enhancer trap, which originated as an important application of P-element vectors (p. 219). The
method employs a lacZ reporter construct, in which
the reporter gene is transcribed from a minimal
promoter, such as a TATA box. Expression from the
promoter is weak, because the promoter lacks an
enhancer to stimulate its transcriptional activity. Pelement-mediated transposition is used to transpose
the construct into many different genomic positions
in separate fly lines. In some flies, by chance, the
construct is transposed to a position where it comes
under the influence of an enhancer that activates
transcription from the weak promoter (Fig. 13.13).
It is often found in practice that, when using a histochemical stain for β-galactosidase activity, the pattern of expression shows cell specificity. Sometimes
the pattern of expression is remarkably refined and
detailed (O’Kane & Gehring 1987). The pattern of
lacZ expression is assumed to reflect the cell-type
specificity of the enhancer. Presumably, an endogenous gene located within range of the enhancer’s
effect has the same pattern of expression as the
reporter. This assumption is known to be valid in
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Randomly transposed
P-element having a weak
promoter driving lacZ
transcription
Enhancer action
lacZ
Genome
Enhancer
Promoter
Fig. 13.13 Enhancer trapping. A reporter construct consists
of lacZ linked to a weak promoter, which requires an enhancer
for significant transcriptional activity. P-element-mediated
transposition is used to insert the construct at random sites
in the fly genome. When the promoter is inserted within
the active range of an enhancer, expression of lacZ can
be detected.
some cases, but the reporter activity does not always
exactly match that of an endogenous gene.
Since enhancers are often found a considerable
distance from the gene they activate, the enhancer
trap cannot be used to directly clone genes by
tagging, although it can be used as the basis for a
chromosome walk to identify the gene of interest.
Enhancer-trap lines have been used to identify and
clone novel Drosophila genes, but the cell-type-specific
expression that is revealed can also be harnessed in
other ways. For example, instead of driving expression of a lacZ reporter, the principle could be applied
to the expression of a toxic gene (such as ricin or
Enhancer trap line
E
diphtheria toxin), leading to cell death and thus
ablation of specific cell lineages in the fly. This has
enormous potential, e.g. in studies of the development of the nervous system (O’Kane & Moffat 1992).
In order to facilitate the use of enhancer trapping
as a general method for driving cell-specific expression, a modified strategy has been developed. This
depends upon the ability of the yeast transcription
factor GAL4 to activate transgenes containing its
recognition site in the heterologous environment
of the fly. The enhancer-trap principle shown in
Fig. 13.13 is modified so that the lacZ reporter gene is
replaced by the coding region for GAL4 (Fig. 13.14).
In such flies, GAL4 is now expressed in the pattern
dictated by a local enhancer, although this cannot
be seen. The pattern of GAL4 expression can be
revealed by crossing the enhancer-trap line to flies
carrying a reporter transgene in which the lacZ gene
is coupled to a promoter containing GAL4-binding
sites. The beauty of this system is that a bank of fly
stocks with different trapped enhancers can be built
up, each with a defined pattern of GAL4 expression.
Once the patterns of GAL4 expression are known,
crosses can be performed to introduce chromosomes
containing constructs in which any desired gene is
coupled to a GAL4-dependent promoter, giving a
particular cell-specific pattern of expression.
UAS-lacZ line
P
GAL 4
E
UAS
P
GAL 4
GAL 4
UAS
lacZ
LacZ
Fig. 13.14 Second-generation
enhancer trap, in which the yeast gene
for the GAL4 transcription factor is
activated by the enhancer. This can
be crossed to a responder line in which
lacZ is driven by a GAL4-dependent
promoter.
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Enhancer traps in other species
Simple lacZ enhancer traps based on the original
Drosophila model have also been used in mice (Allen
et al. 1988, Kothary et al. 1988). The constructs
were introduced into eggs by pronuclear microinjection and the resulting transgenic embryos could be
screened for β-galactosidase activity. One disadvantage of this strategy is that, unlike P-element insertion, gene transfer by pronuclear microinjection
tends to generate complex, multicopy, transgene
arrays, often combined with major structural rearrangements of the genome. These factors complicate
the interpretation of reporter expression patterns
and hamper efforts to recover the corresponding
genes. The use of ES cells and alternative transformation techniques have addressed these problems.
ES cells can be transformed by electroporation,
resulting in simpler transgenic loci with a lower
transgene copy number (Gossler et al. 1989), or
by retroviral vectors, which generate single-copy
insertions (Friedrich & Soriano 1991, Von Melchner
et al. 1992). Enhancer-trap vectors have also been
developed for plants, using T-DNA insertions carrying a gusA gene driven by a weak promoter adjacent
to the right border repeat (Goldsbrough & Bevan
1991, Topping et al. 1991). The activity of endogenous enhancers can be determined by screening the plants for GUS activity. The use of lacZ
and gusA as histological reporters is discussed in
Box 13.1.
Gene traps and functional genomics
Although enhancer traps have been widely used in
animals and plants, there have been relatively few
reports of successful gene-cloning efforts other than
in Drosophila. Much more use has been made of an
alternative type of entrapment construct, the gene
trap, which is activated only upon direct integration
into a functional gene. Depending on the site of
integration, the insertion may or may not generate
a mutant phenotype, but in either case the gene
can be directly cloned by virtue of its unique insertional tag.
The first entrapment vectors in plants were gene
traps that employed a promoterless selectable marker
providing resistance to the antibiotic neomycin, in
271
an attempt to directly select for T-DNA insertions
into functional genes (Andre et al. 1986, Teeri et al.
1986). A problem with this approach was that
selection favoured plants with multiple copies of
abnormal T-DNA inserts (Koncz et al. 1989). A
second selectable marker was therefore incorporated into the vector, allowing transformed plants
to be selected for hygromycin resistance, while the
trapped neomycin-resistance marker was used as a
reporter gene to confirm integration into functional
genes. Replacing the neomycin-resistance marker
with gusA provided the increased versatility of direct
visual screening of gene-trap lines, and vectors of
this type, based on transposons or T-DNA, have been
widely used (Federoff & Smith 1993, Lindsey et al.
1993). An alternative type of gene trap incorporates
a splice acceptor site, in addition to the promoterless
reporter gene, thus making reporter-gene expression dependent on the formation of a transcriptional
fusion with the trapped gene (Sundaresan et al.
1995). Promoter-trap and splice-trap vectors are
compared in Fig. 13.15.
In mice, gene-trap technology has become very
sophisticated and can be used, in ES cells, as an
excellent high-throughput strategy to identify and
characterize new genes. The technique was developed in parallel in a number of laboratories, predominantly based on the splice-trap principle (Gossler
et al. 1989, Friedrich & Soriano 1991, Skarnes et al.
1992, Wurst et al. 1995).
Several clever strategies have been employed to
address potential pitfalls of the technique and improve
the versatility of gene-trap vectors. For example, in
order to select ES cells with productive gene-trap
insertions, the selectable marker neo has been fused
to the reporter gene lacZ to generate a hybrid marker
called β-geo (Gossler et al. 1989). A potential disadvantage of the gene-trap strategy is that only inframe insertions will generate a functional reporter
protein. This has been addressed using an internal
ribosome entry site (Box 13.3), so that translation of
the reporter gene occurs independently of the transcript in which it is embedded (Skarnes et al. 1995,
Chowdhury et al. 1997). The possibility that transiently expressed genes will be overlooked in largescale screens has been addressed, using a binary
system in which Cre recombinase is expressed in the
gene-trap construct and activates the lacZ reporter
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Enhancer trap
Gene trap
P ATG
E
SA ATG
P
ATG
SD
SA
ATG
Promoter trap
E
Enhancer
P
Promoter
Exon
Intron
Reporter gene
gene, using the RAGE strategy discussed earlier in the
chapter (p. 257). Thus, β-galactosidase is expressed
constitutively in all cells where Cre was transiently
active (Thorey et al. 1998). In other vectors, the reporter gene incorporates a nuclear targeting sequence,
which carries the enzyme into the nucleus and
increases the concentration of the signal (Takeuchi
et al. 1995). However, relying on endogenous tar-
Fig. 13.15 Comparison of the structure
and target-integration positions of three
different types of entrapment vector – the
enhancer trap, a typical gene trap and a
promoter-dependent gene trap.
geting signals provided by the interrupted gene can
allow visual screening for proteins that function in
specific cell compartments (e.g. Tate et al. 1998).
The applicability of gene trapping to highthroughput functional genomics has been demonstrated by Wiles and colleagues, who were able to
generate a library of over 12 000 gene-trapped ES
cell lines, using retroviral gene-trap vectors (Wiles
Box 13.3 Internal ribosome entry sites
The translation of most eukaryotic mRNAs is
dependent on the 5′ cap, with which the ribosome
associates before scanning for the initiation codon.
However, certain RNA viruses and a few endogenous
mRNAs contain an internal ribosome entry site (IRES),
which allows cap-independent translation (reviewed
by Mountford & Smith 1995).
An IRES can be very useful in expression vectors
because it allows multiple transgenes to be expressed
on a single transcript. One way in which this
can be exploited is to increase the efficiency of
cotransformation. In traditional plasmid-based
cotransformation techniques, the transgene and a
selectable marker are transcribed independently;
therefore many selected cells express the marker
but not the transgene. However, if both genes are
expressed as a single, dicistronic mRNA, then
selection for the marker necessarily identifies cells
in which the non-selected transgene is also expressed
(Kaufman et al. 1991). Cotransformation with
retroviral vectors is also much more efficient if a
dicistronic expression system is used in preference
to dual-promoter or alternative-splicing vectors, since,
in the latter cases, the non-selected gene is often not
expressed (e.g. Ghattas et al. 1991).
Linking an IRES to a reporter gene also has
numerous applications. For example, most transgenes
do not confer a convenient phenotype that can be
used to track their expression. However, by placing
an IRES-controlled reporter gene downstream of the
transgene, the expression pattern of the transgene
can be established very easily. In this context, IRES
elements are particularly useful where transcriptional
fusions are generated, because this removes any
dependence on in-frame insertions to generate a
functional protein. This is important in gene-trap
vectors, where the position of integration cannot
be controlled (Skarnes et al. 1995; see p. 271).
POGC13 9/11/2001 11:13 AM Page 273
Advances in transgenic technology
SA
IRES
lacZ/neo
polyA
P
273
pur
SD
Fig. 13.16 A highly versatile gene-trap vector designed for high-throughput screening. See text for details.
et al. 2000). Such a library of clones can be grown,
stored frozen in microtitre plates and systematically
analysed to build up a molecular library of flanking
sequences. In order to characterize the expression
patterns of the trapped genes, it is possible to undertake large-scale screens of mouse embryos or particular tissues of adult mice produced from the ES
cells, but prescreening is also possible by exploiting
the ability of ES cells to differentiate along different
pathways in specific culture media (e.g. Reddy et al.
1992, Baker et al. 1997, Yang et al. 1997). It is also
possible to identify ES cells in which particular
functional subsets of genes are trapped, e.g. secreted
proteins (see below). An elegant gene-trap vector
suitable for high-throughput screening has been
designed by Zambrowicz et al. (1998) (Fig. 13.16).
There are two expression cassettes in this vector:
the β-geo hybrid marker downstream of a splice
acceptor and internal ribosome entry site, and the
selectable marker pur, conferring resistance to puromycin, under the control of a constitutive promoter
but lacking a polyadenylation signal, upstream of
a splice donor site. Using this vector, expression of
the reporter is dependent on the formation of a
transcriptional fusion with the trapped gene, but,
since the selectable marker is driven by its own
promoter but dependent on the structure of the gene
for correct processing, selection for insertions is not
biased in favour of genes that are actively expressed
in ES cells.
Recently, a Xenopus gene trap has also been
developed, based on the transgenesis procedure of
Kroll and Amaya (1996) (p. 216). In this system, the
reporter gene for green fluorescent protein is used
(Box 13.1), allowing reporter expression patterns
to be analysed in living tadpoles (Bronchain et al.
1999). This system could become a powerful tool for
functional genomics in Xenopus because hundreds
or thousands of embryos can be generated in a single
day and screened for gene-trap events in real time.
Function-specific trapping
The design of gene-trap constructs can be modified
to select for particular classes of genes. For example,
Skarnes et al. (1995) describe a construct in which
the β-geo marker is expressed as a fusion to the
transmembrane domain of the CD4 type I protein. If
this inserts into a gene encoding a secreted product,
the resulting fusion protein contains a signal peptide
and is inserted into the membrane of the ER in the
correct orientation to maintain β-galactosidase
activity. However, if the construct inserts into a
different type of gene, the fusion product is inserted
into the ER membrane in the opposite orientation
and β-galactosidase activity is lost.
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CHAPTER 14
Applications of recombinant
DNA technology
Introduction
Biotechnology is not new. The making of beer, wine,
bread, yoghurt and cheese was practised by ancient
civilizations, such as the Babylonians, the Romans
and the Chinese. Much, much later came vaccines,
the production of basic chemicals (e.g. glycerol,
citric acid, lactic acid) and the development of antibiotics. In each of these examples, existing properties
of microorganisms were exploited. For example,
Penicillium species naturally make penicillin. What
the scientists have done is to increase the yield of
penicillin by repeated rounds of mutation and
selection, coupled with optimization of the growth
medium. Similarly, sexual crosses between related
plant species have created high-yielding and diseaseresistant varieties of cereals. These improved cereals
represent new combinations of genes and alleles
already existing in wild strains.
With the development of gene manipulation techniques in the 1970s, there was a major paradigm
shift. For the first time microorganisms could be
made to synthesize compounds that they had never
synthesized before, e.g. insulin production in E. coli
( Johnson 1983). Soon all sorts of commercially or
therapeutically useful proteins were being made in
bacteria, principally E. coli, and thus the modern
biotechnology industry was born. As the techniques
developed for manipulating genes in bacteria were
extended to plants and animals there was a concomitant expansion of the biotechnology industry
to exploit the new opportunities being provided.
Today there are many different facets to the commercial exploitation of gene manipulation techniques as shown in Fig. 14.1. Rather than discuss
all these topics in detail, for that would take a book
in itself, we have chosen to focus on six interdisciplinary themes that reflect both the successes
achieved to date and the likely successes in the next
decade.
Theme 1: Nucleic acid sequences
as diagnostic tools
Introduction to theme 1
Nucleic acid sequences can be used diagnostically in
two different ways. The first is to determine whether
a particular, relatively long sequence is present in or
absent from a test sample. A good example of such
an application is the diagnosis of infectious disease.
By choosing appropriate probes, one can ascertain
in a single step which, if any, microorganisms are
present in a sample. Alternatively, a search could be
made for the presence of known antibiotic-resistance
determinants so that an appropriate therapeutic
regime can be instituted. In the second way in which
sequences are used diagnostically, the objective is to
determine the similarity of sequences from different
individuals. Good examples of this approach are
prenatal diagnosis of genetic disease and forensic
profiling (‘DNA fingerprinting’).
Detection of sequences at the gross level
Imagine that a seriously ill individual has a disorder
of the gastrointestinal tract. A likely cause is a
microbial infection and there are a number of candidate organisms (Table 14.1). The question is, which
organism is present and to which antibiotics is it
susceptible? The sooner one has an answer to these
questions, the sooner effective therapy can begin.
Traditionally, in such a case, a stool specimen would
be cultured on a variety of different media and would
be examined microscopically and tested with various immunological reagents. A simpler approach
is to test the sample with a battery of probes and
determine which, if any, hybridize in a simple dotblot assay. With such a simple format it is possible to
vary the stringency of the hybridization reaction to
accommodate any sequence differences that might
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Applications of recombinant DNA technology
275
Transgenic
animals
Nucleic
acids
Therapeutic proteins
Anti-sense
Improved farm
animals
Gene therapy/repair
Diagnostic probes
‘Pharming’
Disease models
Vaccines
Recombinant
DNA
technology
Vaccines
Vaccines
Small molecules
Small molecules
Therapeutic proteins
Therapeutic proteins
Improved plants
Enzymes
Biopolymers
Transgenic
plants
Recombinant
microbes
Fig. 14.1 The different ways that recombinant DNA technology has been exploited.
Table 14.1 Pathogens causing infection of the
gastrointestinal tract.
Bacteria
Salmonella spp.
Shigella spp.
Yersinia enterocolitica
Enterotoxigenic E. coli
Clostridium difficile
Clostridium welchii
Campylobacter spp.
Aeromonas spp.
Vibrio spp.
Viruses
Rotaviruses
Enteroviruses
Enteric adenoviruses
Protozoa
Entamoeba histolytica
Giardia Iamblia
exist between the probe and the target. The downside of this approach is that if one wishes to test the
sample with 10 different probes, then 10 different
dot blots are required; otherwise there is no way of
determining which probe has bound to the target.
Another disadvantage of this approach is that
sufficient target DNA must be present in the sample
to enable a signal to be detected on hybridization.
Both of these problems can be overcome by using the
polymerase chain reaction (PCR).
In a traditional dot-blot assay, the sample DNA is
immobilized on a membrane and hybridized with a
selection of labelled probes. An alternative is a
‘reverse dot blot’, where the probes are immobilized
on the membrane and hybridized with the sample.
In this way, only one hybridization step is required,
because each probe occupies a unique position on
the membrane (Fig. 14.2). For this approach to
work, the sample DNA needs to be labelled and this
can be achieved using the PCR. This has the added
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CHAPTER 14
Conventional dot blot
Membrane
Test DNA
Probe
A
Test DNA
Probe
B
Test DNA
Probe
C
Test DNA
Probe
D
Test DNA
Probe
E
Reverse dot blot
Probe A
Probe B
Probe C
Probe D
Labelled
test DNA
benefit of amplifying the sample, thereby minimizing
the amount of target DNA required. The downside
of this approach is that the amplification step needs
to be done with multiple pairs of primers (multiplex
PCR), one for each target sequence. Although
multiplex PCR is now an established method, considerable optimization is needed for each application
(Elnifro et al. 2000). The major problems encountered are poor sensitivity or specificity and/or preferential amplification of certain specific targets.
The presence of more than one primer pair in the
reaction increases the chance of obtaining spurious
amplification products because of the formation of
primer dimers. Such non-specific products may be
amplified more efficiently than the desired target.
Clearly, the design of each primer pair is crucial to
avoiding this problem. Another important feature is
that all primer pairs should enable similar amplification efficiencies for their respective targets.
Probe E
Fig. 14.2 Comparison of conventional
dot blot assay with reverse dot blot
method. Note that in the latter there
is only a single hybridization reaction,
regardless of the number of probes
used, whereas in the former each
probe has to be tested in a separate
hybridization step.
Despite the difficulties noted above, multiplex PCR
has been used successfully in the diagnosis of infectious diseases. For example, Heredia et al. (1996)
have used the method to simultaneously examine
blood for the presence of HIV-1, HIV-2, human
T-cell lymphotropic virus (HTLV)-I and HTLV-II.
Similarly, Grondahl et al. (1999) have used the
method to identify which of nine different organisms
is responsible for respiratory infections. Once the
clinical microbiologist knows the identity of a microorganism in a specimen, he/she can select those
antibiotics that might be effective, provided that
the organism in question does not carry multiple
drug-resistance determinants. The presence of these
determinants can also be established using multiplex PCR. Alternatively, if a virus causes the infection, one can monitor the progress of the infection by
using quantitative PCR (see p. 23). This approach
has been used to monitor cytomegalovirus infec-
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Applications of recombinant DNA technology
tions in kidney-transplant patients (Caballero et al.
1997). It is worth noting that the development
of multiplex PCR technology is being facilitated by
the rapid progress in the sequencing of microbial
genomes (for an up-to-date list, see http//igweb.integratedgenomics.com/GOLD/), since the data generated enable species-specific genes or sequences to be
identified.
Comparative sequence analysis: singlenucleotide polymorphisms (SNPs)
In prenatal diagnosis of genetic disorders, there is
a need to determine which alleles of a particular
locus are being carried by the fetus, i.e. is the fetus
homozygous for the normal or the deleterious allele
or is it heterozygous? In forensic DNA profiling the
requirement is to match DNA from the perpetrator
of a crime with that of a suspect. In each case, a
definitive answer could be obtained by sequencing
the relevant samples of DNA. While this is possible, it
is not practicable for mass screening. An alternative
could be to detect hybridization of specific probes.
This has been done for the detection of sickle-cell
anaemia by Conner et al. (1983). They synthesized
Marker
DNA
Fig. 14.3 Schematic representation
of the use of oligonucleotide probes
to detect the normal α1-antitrypsin
gene (M) and its Z variant. Human
DNA obtained from normal (MM),
heterozygous (MZ) and homozygous
variant (ZZ) subjects is digested
with a restriction endonuclease,
electrophoresed and fragments
Southern blotted on to a nitrocellulose
membrane. The patterns shown were
obtained on autoradiography of the
filter following hybridization with
either the normal (M-specific) or
variant (Z-specific) probe.
MM
277
two 19-mer oligonucleotides, one of which was
complementary to the amino-terminal region of the
normal β-globin (βA) gene and the other of which
was complementary to the sickle-cell β-globin gene
(βS). These oligonucleotides were radiolabelled and
used to probe Southern blots. Under appropriate
conditions, the probes could distinguish the normal
and mutant alleles. The DNA from normal homozygotes only hybridized with the βA probe and DNA
from sickle-cell homozygotes only hybridized with
the βS probe. DNA of heterozygotes hybridized with
both probes. These experiments, therefore, showed
that oligonucleotide hybridization probes can discriminate between a fully complementary DNA and
one containing a single mismatched base. Similar
results have been obtained (Fig. 14.3) with a point
mutation in the α-antitrypsin gene, which is implicated in pulmonary emphysema (Cox et al. 1985).
The single-base changes that occur in the two
clinical examples just quoted are examples of singlenucleotide polymorphisms (SNPs, pronounced
‘snips’). Many such polymorphisms occur throughout an entire genome and in humans the frequency
is about once every 1000 bases. Their distribution is
not entirely random. SNPs that alter amino acid
MZ
ZZ
M-specific probe
(C TTT CTC GTC GAT GGT CAG)
Marker
DNA
MM
MZ
ZZ
Z-specific probe
(C TTT CTT GTC GAT GGT CAG)
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278
CHAPTER 14
MstII
MstII
*
AA
Amn
AS
SS
(b)
(a)
MstII
β-globin gene
*
Pro Glu Glu
CCTGAGGAG
5
6
7
CCTGTGGAG
Pro Val Glu
Base sequence in
normal (β A) gene
Base sequence in
mutant (sickle, β S ) gene
βA
βS
Fig. 14.4 Antenatal detection of sickle-cell genes. Normal individuals are homozygous for the βA allele, while sufferers from
sickle-cell anaemia are homozygous for the βS allele. Heterozygous individuals have the genotype βAβS. In sickle-cell anaemia,
the 6th amino acid of β-globin is changed from glutamate to valine. (a) Location of recognition sequences for restriction
endonuclease MstII in and around the β-globin gene. The change of A → T in codon 6 of the β-globin gene destroys the
recognition site (CCTGAGG) for MstII as indicated by the asterisk. (b) Electrophoretic separation of MstII-generated fragments
of human control DNAs (AA, AS, SS) and DNA from amniocytes (Amn). After Southern blotting and probing with a cloned
β-globin gene, the normal gene and the sickle gene can be clearly distinguished. Examination of the pattern for the amniocyte
DNA indicates that the fetus has the genotype βAβS, i.e. it is heterozygous.
sequences occur much less frequently than silent
substitutions and SNPs in non-coding regions (Cargill
et al. 1999). However, they are stably inherited.
In some instances, these polymorphisms result in
the creation or elimination of a restriction-enzyme
site and this can be used diagnostically. A classical
example is sickle-cell anaemia (Fig. 14.4), where
the mutation from GAG to GTG eliminates restriction sites for the enzymes DdeI (CTNAG) and MstII
(CCTNAGG) (Chang & Kan 1981, Orkin et al. 1982).
The mutation has been detected by digesting mutant
and normal DNA with the restriction enzyme and
performing a Southern-blot hybridization with a
cloned β-globin DNA probe.
Many of the SNPs that cause genetic diseases
do not lie within a restriction-enzyme site, as is the
case in sickle-cell anaemia. However, the restriction
fragment length poymorphisms (RFLPs) caused by
other SNPs can be used diagnostically, as shown
in Fig. 14.5. In this case there is a close linkage
between a polymorphic restriction site and the locus
of interest and this can be used to trace the inheritance of the gene. When this approach was first
developed, a major limitation was the availability
of suitable polymorphic markers. Following the
sequencing of the entire human genome, an encyclopedia of SNPs is being created (currently 1.4
million) and this will greatly facilitate association
studies. Indeed, this approach is now being used to
match patients with appropriate drugs (see p. 292).
Again, when this approach was developed, RFLPs
were detected by genomic Southern blotting. This
laborious step can be bypassed by use of the PCR.
Enough DNA can be synthesized in the PCR reaction
so that after digestion with the restriction enzyme
and electrophoresis, the DNA bands are directly
visible following staining with ethidium bromide.
Another advantage of PCR is that the gel step can be
omitted altogether and the SNPs detected directly,
using microarrays (‘DNA chips’) (p. 116).
POGC14 9/11/2001 11:11 AM Page 279
Applications of recombinant DNA technology
10 kb
P
M
N
M
A
10 kb
7 kb
7 kb
Gene map of
parents
Chromosomes of parents
Mother +–
Father
–
+
10 kb
Brother ++
Fetus
7 kb
+
+
Family studied by gene mapping
Gene map of
brother and fetus
Fig. 14.5 An example of prenatal diagnosis using restriction
fragment length polymorphism (RFLP) linkage analysis. The
parents are both carriers for a deleterious gene (A): one of
their chromosomes carries this determinant, the other its
normal allele (N). One of the parental chromosomes carries
a polymorphic restriction enzyme site P, which is close
enough to A or N for them not to be separated in successive
generations. On the chromosome which does not contain this
site (–), a particular restriction enzyme cuts out a piece of DNA
10 kb long which contains another locus (M), for which we
have a radioactive probe. On the chromosome containing the
polymorphism (+), a single base change produces a new site
and hence the DNA fragment containing locus M is now only
7 kb. On gene mapping of the parents’ DNA using probe M,
we see two bands representing either the + or − chromosomes.
A previously born child had received the deleterious gene
A from both parents and on mapping we find that it has the
++ chromosome arrangement, i.e. only a single 7 kb band.
Hence the mutation must be on the + chromosome in both
parents. To identify the disease in a fetus in subsequent
pregnancies, we shall be looking for an identical pattern,
i.e. the 7 kb band only. (Reproduced courtesy of Professor
D. Weatherall and Oxford University Press.)
Variable number tardem repeat
(VNTR) polymorphisms
In higher eukaryotes, genes and their associated
introns occupy only a small proportion of the
genome. The intergenic DNA, which makes up the
majority of the genome, is composed of a mixture of
279
unique sequences and repetitive sequences. Many of
the repetitive DNA sequence elements are arranged
in tandem and are known as satellite DNA. Three
types of satellite DNA can be distinguished on the
basis of the level of repetition and the repeat-unit
length (Table 14.2). Not only is satellite DNA dispersed throughout the genome, it is highly variable
and provides a valuable tool for genetic individualization. An example of this is shown schematically
in Fig. 14.6. In this case the RFLPs detected are due
to variations in the number of repeat units (VNTR
polymorphisms) between restriction sites, rather
than changes in the location of the restriction sites,
as discussed in the previous section.
VNTR polymorphisms are of importance to
clinical geneticists because a number of important
hereditary diseases are associated with alterations
in the degree of repetition of microsatellites (for
reviews see Bowater & Wells 2000, Gutekunst et al.
2000, Usdin & Grabczyck 2000). Probably the best
example of such a disease is Huntington’s chorea,
which is caused by an expansion of a CAG trinucleotide repeat in exon 1 of the gene coding for a
protein of unknown function, which has been
named huntingtin. Expansion beyond 40 repeat
units correlates with the onset and progression of
the disease (for review see Reddy et al. 1999).
Forensic applications of VNTRs
The existence of VNTR polymorphisms is of great
utility in paternity testing and criminal investigations,
since they allow ready comparison of DNA samples
in the absence of detailed genetic information by the
generation of a DNA profile or fingerprint. In principle, a multilocus DNA fingerprint can be generated
either by the simultaneous application of several
probes, each one specific for a particular locus, or by
applying a single DNA probe that simultaneously
detects several loci. When DNA profiling was first
developed (Jeffreys et al. 1985a), multilocus probes
were used and these were derived from a tandemly
repeated sequence within an intron of the myoglobin gene (Fig. 14.7). These probes can hybridize
to other autosomal loci – hence their utility. The
first criminal court case to use DNA fingerprinting
was in Bristol, UK, in 1987, when a link was shown
between a burglary and a rape. In the following
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CHAPTER 14
Table 14.2 Classification of satellite
DNA.
Type of repeat
Degree of repetition
per locus
Satellite
103–107
Minisatellite
10–103
Microsatellite
10–102
H1
Number of loci
One to two
per chromosome
Thousands per
genome
Up to 105
per genome
Repeat-unit
length (bp)
1000–3000
9–100
1–6
H1
Core sequence
Individual
A
Probe 33.6
Probe 33.15
Probe 33.5
Individual
B
A
GGAGGTGGGCAGGAGG
[(A G G G C T G G A G G )3]18
(A G A G G T G G G C A G G T G G)29
C
(G G G A G T G G G C A G G A G G)14
Fig. 14.7 Probes used for DNA fingerprinting.
Individual
C
Individual
A
Individual
B
Individual
C
Fig. 14.6 Restriction fragment length polymorphisms
caused by a variable number of tandem repeats between
the two HinfI restriction sites. The upper part of the diagram
shows the DNA structure for three different individuals.
The lower part of the diagram shows the pattern obtained on
electrophoresis of HinfI cut DNA from the three individuals
after hybridization with a probe complementary to the
sequence shown in pink.
year, DNA-fingerprinting evidence was used in the
USA. It is worth noting that DNA evidence has been
used to prove innocence as well as guilt (Gill &
Werrett 1987).
Multilocus probes can also be used to prove or
disprove paternity and a unique example, which
was part of an immigration test case, is shown in
Box 14.1. The technique also has application in
many other areas, such as pedigree analysis in cats
and dogs ( Jeffreys & Morton 1987), confirming
cell-line authenticity in animal cell cultures (Devor
et al. 1988, Stacey et al. 1992) and monitoring the
behaviour and breeding success of bird populations
(Burke & Bruford 1987).
In criminal cases, a major disadvantage of multilocus probes is the complexity of the DNA fingerprint
provided. Showing innocence is easy, but proving
identity is fraught with problems. The issue boils
down to calculations of the probability that two
profiles match by chance as opposed to having come
from the same person (Lewontin & Hartl 1991). For
this reason forensic scientists have moved to the use
of single-locus probes and an example is shown in
Fig. 14.8. The latest variation of the technique,
introduced in the UK in 1999, targets 10 distinct
loci, and the likelihood of two people sharing the
same profile is less than one in a billion (thousand
million). Chance matches are even less likely in the
USA, where the FBI routinely examines 13 VNTR
loci. Another advantage of single-locus probes is
POGC14 9/11/2001 11:11 AM Page 281
Applications of recombinant DNA technology
281
Box 14.1 Use of DNA fingerprinting in an immigration test case
In 1984, a Ghanaian boy was refused entry into
Britain because the immigration authorities were
not satisfied that the woman claiming him as her son
was in fact his mother. Analysis of serum proteins and
erythrocyte antigens and enzymes showed that the
alleged mother and son were related but could not
determine whether the woman was the boy’s mother
or aunt. To complicate matters, the father was not
available for analysis nor was the mother certain
of the boy’s paternity. DNA fingerprints from blood
samples taken from the mother and three children
who were undisputedly hers, as well as the alleged
son, were prepared by Southern blot hybridization to
two of the mini-satellite probes shown in Fig. B14.1.
Although the father was absent, most of his DNA
fingerprint could be reconstructed from paternalspecific DNA fragments present in at least one of
the three undisputed siblings but absent from the
mother. The DNA fingerprint of the alleged son
contained 61 scorable fragments, all of which
were present in the mother and/or at least one of the
siblings. Analysis of the data showed the following.
• The probability that either the mother or the
father by chance possess all 61 of the alleged son’s
bands is 7 × 10−22. Clearly the alleged son is part
of the family.
• There were 25 maternal-specific fragments in
the 61 identified in the alleged son and the chance
probability of this is 2 × 10−5. Thus the mother and
alleged son are related.
• If the alleged mother of the boy in question is in
fact a maternal aunt, the chance of her sharing the
25 maternal-specific fragments with her sister is
6 × 10−6.
When presented with the above data (Jeffreys et al.
1985b), as well as results from conventional marker
analysis, the immigration authorities allowed the boy
residence in Britain. In a similar kind of investigation,
a man originally charged with murder was shown to
be innocent (Gill & Werrett 1987).
U
kb
20
10
8
Fig. B14.1 DNA fingerprints of a Ghanaian
family involved in an immigration dispute.
Fingerprints of blood DNA are shown for
the mother (M), the boy in dispute (X), his
brother (B), sisters (S1, S2) and an unrelated
individual (U). Fragments present in the
mother’s (M) DNA are indicated by a
short horizontal line (to the right of each
fingerprint); paternal fragments absent
from M but present in at least one of the
undisputed siblings (B, S1, S2) are marked
with a long line. Maternal and paternal
fragments transmitted to X are shown with
a dot. (Photo courtesy of Dr A. Jeffreys and
the editor of Nature.)
6
4
2
M
X
B
S1
S2
M
X
B
S1
S2
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C
CHAPTER 14
A
D
Anorak Swab
D
A
worn by a bank robber, a cigarette-butt discarded
at the scene of a crime or the back of a stamp on an
envelope used to send a ‘poison-pen’ or blackmail
letter.
C
Historical genetics
4 kb
Fig. 14.8 Use of a single locus probe to determine the
identity of a rapist. Semen was extracted from an anorak
and a vaginal swab. The victim’s profile is in track D and
that of two suspects in tracks A and C. The profile matches
individual A. (Photo courtesy of Dr. P. Gill.)
that it is possible to convert the DNA profile into a
numerical format. This enables a database to be
established and all new profiles can be matched to
that database.
Detection of VNTR polymorphisms requires that
an adequate amount of DNA be present in the test
sample. This is not a problem in paternity disputes
but can be an issue in forensic testing. With the
advent of single-locus probes, the amount of DNA
required is much less of an issue, since the test loci
in the sample can be amplified by PCR. As a result,
it now is possible to type DNA from a face-mask
Just as multilocus probes have been used for many
applications other than in crime testing, so too have
single-locus probes. A good example is the determination of the parentage of grapevines used for wine
making. Grapevines are propagated vegetatively, so
that individual vines of a cultivar are genetically
identical to each other and to the single original
seedling from which the cultivar originated. Most of
the cultivars in existence in north-eastern France
are centuries old and their provenance was not
known. However, using 17 microsatellite loci,
Bowers et al. (1999) were able to show that 16 of
the common cultivars have genotypes consistent
with their being progeny of a single pair of grape
cultivars that were widespread in the region in the
Middle Ages.
VNTRs can be found in mitochondrial DNA, as
well as nuclear DNA, and these have particular
applications. The reasons for this are threefold. First,
mitochondrial sequences are passed from mother to
child in the egg. Thus, brothers and sisters have
identical mitochondrial DNA. Secondly, the small
size of mitochondrial DNA (16–20 kb) means that
there is less scope for variability, but this is more
than compensated for by the copy number (~10 000
copies per cell). That is, mitochondrial DNA is
naturally amplified. Thirdly, in very old or degraded
specimens, the nuclear DNA may be totally decomposed, but mitochondrial DNA can still be
recovered. For example, mitochondrial-DNA analysis was used to confirm that skeletons found in
Ekaterinburg, Russia, were the remains of the last
tsar and his family (Gill et al. 1994). A similar analysis showed that an individual living in Cheddar
Gorge in the UK was related to a Stone Age individual whose skull was found nearby. Since bones
are more likely than soft tissue to survive in the
event of major accidents that involve fire, mitochondrial DNA analysis will play an increasingly
important role in identifying victims. Indeed, such
an analysis was done in the UK following the 1999
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Paddington train crash, in which one carriage was
completely incinerated.
Just as the mitochondrion is transmitted maternally,
the Y chromosome is transmitted only through male
descendants. Because there is only a single copy of
the Y chromosome in normal diploid cells, recombination between different Y chromosomes does not
occur. Any changes that do occur in the Y chromosome from generation to generation must arise
from DNA rearrangements or by accumulation of
random mutations. That is, the Y chromosome
should be highly conserved. Sykes and Irven (2000)
obtained proof of this. They probed a randomly
ascertained sample of males with the surname ‘Sykes’
with four Y-chromosome microsatellites and found
that half of them had the same Y haplotype. This
suggests that all those with the same haplotype have
a common ancestor, even though conventional
genealogical analysis suggests otherwise.
Theme 2: New drugs and new therapies
for genetic diseases
Introduction to theme 2:
proteins as drugs
One of the earliest commercial applications of
gene-manipulation techniques was the production
in bacteria of human proteins with therapeutic
applications. Not surprisingly, the first such products were recombinant versions of proteins already
used as therapeutics: human growth hormone and
insulin. Prior to the advent of genetic engineering,
human growth hormone was produced from pituitary glands removed from cadavers. Not only did
this limit the supply of the hormone but, in some
cases, it resulted in recipients contracting Creutzfeld–
Jakob syndrome. The recombinant approach resulted
in unlimited supplies of safe material. This safety
aspect has been extended to various clotting factors
that were originally isolated from blood but now
carry the risk of HIV infection. As the methods for
cloning genes became more and more sophisticated,
an increasing number of lymphokines and cytokines
were identified and significant amounts of them
produced for the first time. A number of these were
shown to have therapeutic potential and found their
way into clinical practice (Table 14.3).
283
The first generation of protein drugs were exact
copies of the human molecules but protein engineering is now being used to develop second-generation
molecules with improved properties (see theme 4,
p. 299). More recently, macromodifications have
been made to proteins, as exemplified by the recently
approved drug (Table 14.3) for rheumatoid arthritis, which consists of the tumour necrosis factor
receptor fused to the Fc portion of human IgG1.
Transgenic animals and plants as
bioreactors: ‘pharming’
‘Pharming’ is the play on words that refers to the use
of transgenic animals and plants to produce recombinant therapeutic proteins. As discussed in earlier
chapters, recombinant-protein synthesis in animal
cells has a number of advantages over microbial
expression systems, the most important of which
is the authentic post-translational modifications
that are performed in animal cells. However, largescale culture of animal cells is expensive, due to
the amount of medium and serum required and
the necessity for precise and constant growth conditions. The production of growth hormone in the
serum of transgenic mice (Palmiter et al. 1982a) (see
p. 209) provided the first evidence that recombinant
proteins could be produced, continuously, in the
body fluids of animals. Five years later, several groups
reported the secretion of recombinant proteins in
mouse milk. In each case, this was achieved by joining the transgene to a mammary-specific promoter,
such as that from the casein gene. The first proteins
produced in this way were sheep β-lactoglobulin
(Simons et al. 1987) and human tissue-plasminogen
activator (tPA) (Gordon et al. 1987, Pittius et al.
1988). There have been over 100 such reports since
these early experiments, and a selection is listed in
Table 14.4.
Although proteins can be produced at high concentrations in mouse milk (e.g. 50 ng/ml for tPA),
the system is not ideal, due to the small volume of
milk produced. Therefore, other animals, such as
sheep and goats, have been investigated as possible bioreactors. Such animals not only produce
large volumes of milk, but the regulatory practices
regarding the use of their milk are more acceptable. An early success was Tracy, a transgenic ewe
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Table 14.3 Some recombinant proteins that are used therapeutically.
Year
Product
Clinical indication
1982
1985
1986
Human insulin
Human growth hormone
Hepatitis B vaccine
Interferon-a2a
Tissue plasminogen activator
Erythropoietin
Interferon-g1b
Granulocyte–macrophage-colony-stimulating factor
Granulocyte-colony-stimulating factor
Human interleukin-2
Factor VIII
Human DNase
Glucocerebrosidase
Interferon-b1a
Factor IX
Consensus interferon
Platelet growth factor
Platelet-derived growth factor b
Tumour necrosis factor receptor linked to Fc portion
of human IgG1
Glucagon
Factor VIIa
Diabetes
Pituitary dwarfism
Prevention of hepatitis B infection
Hairy-cell leukaemia
Acute myocardial infarction
Anaemia associated with chronic renal failure
Chronic granulomatous disease
Bone-marrow transplant
Chemotherapy-induced neutropenia
Renal-cell carcinoma
Haemophilia A
Cystic fibrosis
Gaucher’s disease
Multiple sclerosis
Haemophilia B
Chronic HCV infection
Chemotherapy-induced thrombocytopenia
Lower-extremity diabetic ulcers
Rheumatoid arthritis
1987
1989
1990
1991
1992
1993
1994
1996
1997
1998
1999
Hypoglycaemia
Haemophilia
Table 14.4 Some recombinant proteins produced in the secretions of animal bioreactors.
System
Species
Product
Mouse
Rabbit
Sheep
Goat
Sheep b-lactoglobulin
Human tissue-plasminogen activator
Human urokinase
Human growth hormone
Human fibrinogen
Human nerve growth factor
Spider silk
Human erythropoietin
Human a1-antitrypsin
Human tissue-plasminogen activator
Simons et al. 1987
Gordon et al. 1987
Meade et al. 1990
Devinoy et al. 1994
Prunkard et al. 1996
Coulibaly et al. 1999
Karatzas et al. 1999
Massoud et al. 1996
Wright et al. 1991
Ebert et al. 1991
Blood serum
Rabbit
Pig
Human a1-antitrypsin
Recombinant antibodies
Massoud et al. 1991
Lo et al. 1991, Weidle et al. 1991
Urine
Mouse
Human growth hormone
Kerr et al. 1998
Semen
Mouse
Human growth hormone
Dyck et al. 1999
Milk
Reference
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Applications of recombinant DNA technology
producing extremely high levels (30 g/l) of human
α1-antitrypsin (AAT) in her milk (Wright et al. 1991).
Artificially inseminated eggs were microinjected
with a DNA construct containing an AAT gene
fused to a β-lactoglobin promoter. These eggs were
implanted into surrogate mothers, of which 112
gave birth. Four females, including Tracy, and one
male were found to have incorporated intact copies
of the gene and all five developed normally. Over the
lactation period, sheep can produce 250–800 l of
milk, so the production potential is significant.
Using similar protocols, Ebert et al. (1991) have
demonstrated the production of a variant of human
tPA in goat milk. Of 29 offspring, one male and one
female contained the transgene. The transgenic
female underwent two pregnancies and one out of
five offspring was transgenic. Milk collected over her
first lactation contained only a few milligrams of tPA
per litre, but improved expression constructs have
since resulted in an animal generating several
grams per litre of the protein. Recombinant human
antithrombin III, which is used to prevent blood
clots forming in patients that have undergone heartbypass operations, was the first protein expressed
in transgenic animal milk to reach commercial
production, and is currently marketed by Genzyme
Transgenics Corporation.
The production of foreign proteins in secreted
body fluids has the obvious advantage that transgenic animals can be used as a renewable source of
the desirable molecule. In addition to milk, other
production systems have been investigated, including serum (Massoud et al. 1991), semen (Dyck et al.
1999) and urine (Kerr et al. 1998). In each case, an
important consideration is whether the protein is
stable and whether it folds and assembles correctly.
The assembly of complex proteins comprising up to
three separate polypeptides has been demonstrated
in milk, e.g. fibrinogen (Prunkard et al. 1996), collagen ( John et al. 1999) and various immunoglobulins
(e.g. Castilla et al. 1998). Other abundantly secreted
fluids that are likely to be exploited for recombinantprotein expression in the future include the albumen
of hens’ eggs and silkworm cocoons. There has
already been some success with the latter, using
both microinjection (e.g. Nagaraju et al. 1996) and
infection of silkworm larvae with baculovirus
vectors (Tamura et al. 1999, Yamao et al. 1999) (see
285
Chapter 10). The use of animals as bioreactors has
been extensively reviewed (Clark 1998, Rudolph
1999, Wall 1999, Houdebine 2000).
Plants as bioreactors
Plants are a useful alternative to animals for
recombinant-protein production because they are
inexpensive to grow and scale-up from laboratory
testing to commercial production is easy. Therefore,
there is much interest in using plants as production
systems for the synthesis of recombinant proteins
and other speciality chemicals. There is some concern that therapeutic molecules produced in animal
expression systems could be contaminated with
small quantities of endogenous viruses or prions, a
risk factor that is absent from plants. Furthermore,
plants carry out very similar post-translational modification reactions to animal cells, with only minor
differences in glycosylation patterns (CabanesMacheteau et al. 1999). Thus plants are quite
suitable for the production of recombinant human
proteins for therapeutic use.
A selection of therapeutic proteins that have been
expressed in plants is listed in Table 14.5. The first
such report was the expression of human growth
hormone, as a fusion with the Agrobacterium nopaline synthase enzyme, in transgenic tobacco and
sunflower (Barta et al. 1986). Tobacco has been the
most frequently used host for recombinant-protein
expression although edible crops, such as rice, are
now becoming popular, since recombinant proteins
produced in such crops could in principle be administered orally without purification. The expression of
human antibodies in plants has particular relevance
in this context, because the consumption of plant
material containing recombinant antibodies could
provide passive immunity (i.e. immunity brought
about without stimulating the host immune system).
Antibody production in plants was first demonstrated by Hiatt et al. (1989) and During et al.
(1990), who expressed full-size immunoglobulins in
tobacco leaves. Since then, many different types of
antibody have been expressed in plants, predominantly tobacco, including full-size immunoglobulins,
Fab fragments and single-chain Fv fragments (scFvs).
For example, a fully humanized antibody against
herpes simplex virus-2 (HSV-2) has been expressed
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Table 14.5 A selection of pharmaceutical recombinant human proteins expressed in plant systems.
Species
Tobacco, sunflower (plants)
Tobacco, potato (plants)
Tobacco (plants)
Rice (plants)
Tobacco (cell culture)
Tobacco (plants)
Tobacco (cell culture)
Tobacco (root culture)
Rice (cell culture)
Tobacco (seeds)
Tobacco (chloroplasts)
Recombinant human product
Reference
Growth hormone
Serum albumin
Epidermal growth factor
a-Interferon
Erythropoietin
Haemoglobin
Interleukins-2 and 4
Placental alkaline phosphatase
a1-Antitrypsin
Growth hormone
Growth hormone
Barta et al. 1986
Sijmons et al. 1990
Higo et al. 1993
Zhu et al. 1994
Matsumoto et al. 1995
Dieryck et al. 1997
Magnuson et al. 1998
Borisjuk et al. 1999
Terashima et al. 1999
Leite et al. 2000
Staub et al. 2000
in soybean (Zeitlin et al. 1998). Even secretory IgA
(sIgA) antibodies, which have four separate polypeptide components, have been successfully expressed
in plants. This experiment involved the generation
of four separate transgenic tobacco lines, each
expressing a single component, and the sequential
crossing of these lines to generate plants in which
all four transgenes were stacked (Ma et al. 1995).
Plants producing recombinant sIgA against the oral
pathogen Streptococcus mutans have been generated
(Ma et al. 1998), and these plant-derived antibodies
(‘plantibodies’) have recently been commercially
produced as the drug CaroRxTM, marketed by Planet
Biotechnology Inc. A number of other biotechnology
companies are bringing antibody-expressing transgenic plants into commercial production (see Fischer
& Emans 2000).
The impact of genomics
Many of the drugs currently on the market treat the
symptoms of the disease rather than the cause of the
disease. This is analogous to reversing a mutant
phenotype by selecting a mutation at a second
site. Not surprisingly, many of these drugs have
side-effects quite separate from those (e.g. toxicity)
caused by their metabolism. Now that the first drafts
of the human genome sequence are available, it
will be possible to convert disease phenotypes into
nucleotide changes in specific genes (Bailey et al.
2001). These genes will then become targets for new
small-molecule drugs. Already drugs have been
developed based on such genotype–phenotype
correlations. For example, Wettereau et al. (1998)
identified a molecule that normalizes atherogenic
lipoprotein levels caused by a genetic deletion of a
microsomal triglyceride transfer protein.
The identification of genetic changes associated
with particular disease phenotypes offers a number
of novel approaches to the development of therapies.
As well as using such changes as novel targets for
small-molecule drug design, there is an opportunity
to use the techniques described in Chapter 11 to generate animals with the exact same genetic defect and
which can be used as models to test new drug candidates (disease modelling). Furthermore, where drugs
cannot be developed to treat a particular disorder,
there might be an opportunity to correct the disease
by further modification to the genome (gene therapy).
Finally, it is likely that, in the near future, transgenic
animals could be used to provide healthy organs for
humans requiring transplants (xenotransplantation)
(Box 14.2). These topics are discussed in more detail
below.
Transgenic animals as models
of human disease
Mammals have been used as models for human
disease for many years, since they can be exploited to
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287
Box 14.2 Xenotransplantation
Transplantation is widely used to treat organ failure
but there is a shortage of organ donors resulting in
long waiting times and many unnecessary deaths.
In the future, transgenic animals could be used to
supply functional organs to replace failing human
ones. This process is termed xenotransplantation.
There is vigorous debate concerning the ethics of
xenotransplantation but, ethics aside, the technique
remains limited by the phenomenon of hyperacute
rejection, which is caused by the host immune
system. Hyperacute rejection is dependent both on
antibodies raised against the foreign organ and the
activation of the host complement system. In both
cases, the major trigger for rejection appears to be a
disaccharide group (Gal-a(1,3)-Gal) which is present
in pigs but not in primates (Cooper et al. 1994,
Sandrin et al. 1994).
Transgenic strategies have been investigated to
avoid hyperacute rejection, including the expression
of complement-inactivating protein on the cell
surface (Cozzi & White 1995, reviewed by Pearse
et al. 1997), the expression of antibodies against
the disaccharide group (Vanhove et al. 1998)
and attempts to inhibit the expression of a(1,3)
galactosyltransferase, the enzyme that forms this
particular carbohydrate linkage, an enzyme that is
present in pigs but not in primates. In the latter case,
carry out detailed analyses of the molecular basis of
disease and to test newly developed therapeutics
prior to clinical trials in humans. Before the advent
of transgenic animal technology, however, models
of inherited diseases (i.e. diseases with a genetic basis)
were difficult to come by. They could be obtained as
spontaneously occurring mutants, suitable mutant
animals identified in mutagenesis screens and susceptible animal strains obtained by selective breeding.
Gene manipulation now offers a range of alternative
strategies to create specific disease models (see
reviews by Smithies 1993, Bedell et al. 1997, Petters
& Sommer 2000).
Some of the earliest transgenic disease models
were mice predisposed to particular forms of cancer,
because the germ line contained exogenously derived
the simplest strategy would be to knock out the gene
by homologous recombination. Gene targeting has
not yet been achieved in pigs, although the success
of gene targeting/nuclear transfer in sheep (Chapter
11) suggests that knockout pigs could be produced
in the next few years. Alternative procedures include
introducing genes encoding other carbohydratemetabolizing enzymes, so that the Gal-a(1,3)Gal groups are modified into some other less
immunogenic moiety (e.g. Sandrin et al. 1995,
Cohney et al. 1997, Osman et al. 1997). Once
hyperacute rejection has been overcome, there
may be further problems, including delayed rejection,
involving natural killer cells and macrophages (Bach
et al. 1996), and the requirement for T-cell tolerance
(Bracy et al. 1998, Kozlowski et al. 1998, Yang et al.
1998). There are also concerns that endogenous
pig retroviruses could be activated following
transplantation, perhaps even recombining with
human retroviruses to produce potent new hybrids
with unknown properties. Detailed studies have
so far shown no evidence of such a phenomenon (e.g.
Heneine et al. 1998, Patience et al. 1998). For recent
reviews on the prospects of xenotransplantation
(particularly porcine-to-human), the reader should
consult Lambrigts et al. (1998), Sandrin and
McKenzie (1999) and Logan (2000).
oncogenes (e.g. Sinn et al. 1987). This exemplifies
so-called gain-of-function diseases, which are caused
by a dominantly acting allele and can be modelled
simply by adding that allele to the normal genome,
e.g. by microinjection into eggs. Other gain of function diseases that have been modelled in this way
include the Gerstmann–Straussler–Scheinker (GSS)
syndrome, a neurodegenerative disease caused by a
dominantly acting mutated prion protein gene. In
one patient suffering from this disease, a mutation
was identified in codon 102 of the prion protein
gene. Transgenic mice were created carrying this
mutant form of the gene in addition to the wild-type
locus and were shown to develop a similar neurodegenerative pathology to their human counterparts (Hsiao et al. 1990). Other examples of
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gain-of-function disease models include Alzheimer’s
disease, which was modelled by overexpression of the
amyloid precursor protein (Quon et al. 1991), and
the triplet-repeat disorder spinocerebellar ataxia
type 1 (Burright et al. 1995). Simple transgene addition can also be used to model diseases caused by
dominant negative alleles, as recently shown for
the premature ageing disease, Werner’s syndrome
(Wang et al. 2000).
Recessively inherited diseases are generally caused
by loss of function, and these can be modelled by
gene knockout. The earliest report of this strategy
was a mouse model for hypoxanthine-guanine
phosphoribosyltransferase (HPRT) deficiency, generated by disrupting the gene for HRPT (Kuehn et al.
1987). A large number of genes have been modelled
in this way, including those for cystic fibrosis (Dorin
et al. 1992, Snouwaert et al. 1992), fragile-X syndrome (Dutch–Belgian Fragile X Consortium, 1994),
β-thalassaemia (Skow et al. 1983, Ciavattia et al.
1995) and mitochondrial cardiomyopathy (Li et al.
2000). Gene targeting has been widely used to
model human cancers caused by the inactivation
of tumour suppressor genes, such as TP53 and
RB1 (reviewed by Ghebranious & Donehower 1998,
Macleod & Jacks 1999).
While the studies above provide models of singlegene defects in humans, attention is now shifting
towards the modelling of more complex diseases,
which involve multiple genes. This is a challenging
area of research but there have been some encouraging early successes. In many cases, the crossing of
different modified mouse lines has led to interesting
discoveries. For example, undulated mutant mice lack
the gene encoding the transcription factor Pax-1,
and Patch mutant mice are heterozygous for a null
allele of the platelet-derived growth-factor gene.
Hybrid offspring from a mating between these two
strains were shown to model the human birth defect
spina bifida occulta (Helwig et al. 1995). In other
cases, such crosses have pointed the way to possible
novel therapies. For example, transgenic mice overexpressing human α-globin and a mutant form of
the human β-globin gene that promotes polymerization provide good models of sickle-cell anaemia
(Trudel et al. 1991). However, when these mice are
crossed to those ectopically expressing human fetal
haemoglobin in adulthood, the resulting transgenic
hybrids show a remarkable reduction in disease
symptoms (Blouin et al. 2000). Similarly, crossing
transgenic mice overexpressing the anti-apoptotic
protein Bcl-2 to rds mutants, which show inherited
slow retinal degeneration, resulted in hybrid offspring in which retinal degeneration was strikingly
reduced. This indicates that Bcl-2 could possibly
be used in gene therapy to treat the equivalent
human retinal-degeneration syndrome (Nir et al.
2000).
The most complex diseases involve many genes,
and transgenic models would be difficult to create.
However, it is often the case that such diseases can
be reduced to a small number of ‘major genes’ with
severe effects and a larger number of minor genes.
Thus, it has been possible to create mouse models of
Down’s syndrome, which in humans is generally
caused by the presence of three copies of chromosome 21. Trisomy for the equivalent mouse chromosome 16 is a poor model because the two
chromosomes do not contain all the same genes.
However, a critical region for Down’s syndrome has
been identified by studying Down’s patients with
partial deletions of chromosome 21. The generation
of yeast artificial chromosome (YAC) transgenic
mice carrying this essential region provides a useful
model of the disorder (Smith et al. 1997) and has
identified increased dosage of the Dyrk1a (minibrain)
gene as an important component of the learning
defects accompanying the disease. Animal models of
Down’s syndrome have been reviewed (Kola &
Hertzog 1998, Reeves et al. 2001).
Gene transfer to humans – gene therapy
The scope of gene therapy
Gene therapy is any procedure used to treat disease
by modifying the genetic information in the cells of
the patient. In essence, gene therapy is the antithesis
of the disease modelling discussed above. Whereas
disease modelling takes a healthy animal and uses
gene-manipulation techniques to induce a specific
disease, gene therapy takes a diseased animal (or
human) and uses gene-manipulation techniques
in an attempt to correct the disorder and return
the individual to good health. Gene transfer can be
carried out in cultured cells, which are then reintroduced into the patient, or DNA can be transferred
to the patient in vivo, directly or using viral vectors.
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Applications of recombinant DNA technology
X
Mutant (disease) gene – loss of function
D
Mutant (disease) gene – dominant gain of function
289
1 Gene augmentation therapy
Functional
gene
X
X
2 Gene inhibition therapy
Antisense
gene etc.
D
D
3 Gene targeting
Homologous
functional gene
X or
D
4 Assisted killing
Antigen,
cytokine etc.
D
Death
Immune system
stimulated
D
5 Prodrug therapy
Ganciclovir
TK etc
D
Fig. 14.9 Overview of gene-therapy
strategies.
The ex vivo approach can be applied only to certain
tissues, such as bone marrow, in which the cells are
amenable to culture. Gene therapy can be used to
treat diseases caused by mutations in the patient’s
own DNA (inherited disorders, cancers), as well as
infectious diseases, and is particularly valuable in
cases where no conventional treatment exists or
where that treatment is inherently risky. Strategies
include the following (Fig. 14.9):
Death
Confers
sensitivity
D
• Gene-augmentation therapy (GAT), where DNA is
added to the genome with the aim of replacing a
missing gene product.
• Gene targeting to correct mutant alleles.
• Gene-inhibition therapy, using techniques such
as antisense RNA expression or the expression of
intracellular antibodies to treat dominantly acting
diseases.
• The targeted ablation of specific cells.
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Therapeutic gene transfer effectively generates
transgenic human cell clones and, for this reason,
only somatic cells can be used as targets. The prospect
of germ-line transgenesis in humans raises serious
ethical concerns and, with the rapid advances in
technology allowing germ-line transformation and
nuclear transfer in numerous mammals, these
concerns will need to be addressed in the very near
future ( Johnson 1998). As an alternative to permanent gene transfer, transient gene therapy can be
achieved using oligonucleotides, which can disrupt
gene expression at many levels but do not permanently change the genetic material of the cell
(Pollock & Gaken 1995).
The tools and techniques for gene therapy are
essentially similar to those used for gene transfer to
any animal cells. Transfection, direct delivery or
transduction (see Chapter 11) can be used to introduce DNA into cells. Viral vectors are most popular
because of their efficiency of gene transfer in vivo.
However, extreme precautions need to be taken to
ensure the safety of such vectors, avoiding potential
problems, such as the production of infectious viruses
by recombination and the pathological effects of
viral replication. A number of viral vectors have
been developed for gene therapy, including those
based on oncoretroviruses, lentiviruses, adenovirus,
adeno-associated virus, herpes virus and a number
of hybrid vectors combining advantageous elements
of different parental viruses (Robbins et al. 1998,
Reynolds et al. 1999). The risks associated with viral
vectors have promoted research into other delivery
methods, the most popular of which include direct
injection of DNA into tissues (e.g. muscle), the injection of liposome–DNA complexes into the blood and
direct transfer by particle bombardment. Although
inherently much safer than viruses, such procedures
show a generally low efficiency (Scheule & Cheng
1996, Tseng & Huang 1998).
Gene-augmentation therapy for recessive diseases
The first human genetic-engineering experiment was
one of gene marking, rather than gene therapy, and
was designed to demonstrate that an exogenous
gene could be safely transferred into a patient and
that this gene could subsequently be detected in
cells removed from the patient. Both objectives were
met. Tumour-infiltrating lymphocytes (cells that
naturally seek out cancer cells and then kill them by
secreting proteins such as tumour necrosis factor
(TNF)) were isolated from patients with advanced
cancer. The cells were then genetically marked with
a neomycin-resistance gene and injected back into
the same patient (Rosenberg et al. 1990).
The first clinical trial using a therapeutic genetransfer procedure involved a 4-year-old female
patient, Ashanthi DeSilva, suffering from severe combined immune deficiency, resulting from the absence
of the enzyme adenosine deaminase (ADA). This disease fitted many of the ideal criteria for gene-therapy
experimentation. The disease was life-threatening
(therefore making the possibility of unknown treatment-related side-effects ethically acceptable), but
the corresponding gene had been cloned and the
biochemical basis of the disease was understood.
Importantly, since ADA functions in the salvage
pathway of nucleotide biosynthesis (p. 177), cells
in which the genetic lesion had been corrected had
a selective growth advantage over mutant cells,
allowing them to be identified and isolated in vitro.
Conventional treatment for ADA deficiency involves
bone-marrow transplantation from a matching donor.
Essentially the same established procedure could be
used for gene therapy, but the bone-marrow cells
would be derived from the patient herself and would
be genetically modified ex vivo (Fig. 14.10). Cells
from the patient were subjected to leucophaeresis
and mononuclear cells were isolated. These were
grown in culture under conditions that stimulated
T-lymphocyte activation and growth and then
transduced with a retroviral vector carrying a normal ADA gene as well as the neomycin-resistance
gene. Following infusion of these modified cells, both
this patient and a second, who began treatment in
early 1991, showed an improvement in their clinical condition as well as in a battery of in vitro and in
vivo immune-function studies (Anderson 1992).
However, the production of recombinant ADA in
these patients is transient, so each must undergo
regular infusions of recombinant T lymphocytes.
Research is ongoing into procedures for the transformation of bone-marrow stem cells, which would
provide a permanent supply of corrected cells.
Gene-augmentation therapies for a small number
of recessive single-gene diseases are now undergoing
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Applications of recombinant DNA technology
1 Remove bone
marrow from
patient
291
2 Culture
mononuclear cells
ex vivo
3 Transfect
(or transduce) with
functional ADA gene
Fig. 14.10 Procedure for ex vivo gene
therapy, based on the treatment for
ADA deficiency.
clinical trials. We consider cystic fibrosis (CF) as an
example. CF is a disorder that predominantly affects
the lungs, liver and pancreas. The disease is caused
by the loss of a cAMP-regulated membrane-spanning
chloride channel. This results in an electrolyte
imbalance and the accumulation of mucus, often
leading to respiratory failure. CF is a recessive disorder suggesting that the loss of function could
be corrected by introducing a functional copy of
the gene. Indeed, epithelial cells isolated from CF
patients can be restored to normal by transfecting
them with the cloned cystic-fibrosis transmembrane
regulator (CFTR) cDNA. Unlike ADA deficiency, the
cells principally affected by CF cannot be cultured
and returned to the patient, so in vivo delivery strategies must be applied. Targeted delivery of the CFTR
cDNA to affected cells has been achieved using
adenoviral vectors, which have a natural tropism
for the epithelial lining of the respiratory system.
Recombinant viruses carrying the CFTR cDNA have
been introduced into patients using an inhaler
(Zabner et al. 1993, Hay et al. 1995, Knowles et al.
1995). The CFTR cDNA has also been introduced
using liposomes (e.g. Caplen et al. 1995). While such
treatments have resulted in CFTR transgene expression in the nasal epithelium, there were neither consistent changes in chloride transport nor reduction
in the severity of CF symptoms, i.e. they have been
largely ineffective.
5 Return modified
cells to patient
4 Select stable
transformants
Gene-therapy strategies for cancer
Cancer gene therapy was initially an extension of
the early gene-marking experiments. The tumourinfiltrating leucocytes were transformed with a gene
for TNF in addition to the neomycin-resistance gene,
with the aim of improving the efficiency with which
these cells kill tumours by increasing the amount of
TNF they secrete. Although TNF is highly toxic to
humans at levels as low as 10 µg/kg body weight,
there have been no side-effects from the gene therapy and no apparent organ toxicity from secreted
TNF (Hwu et al. 1993). One alternative strategy is
to transform the tumour cells themselves, making
them more susceptible to the immune system through
the expression of cytokines or a foreign antigen.
Another is to transform fibroblasts, which are easier
to grow in culture, and then co-inject these together
with tumour cells to provoke an immune response
against the tumour. A number of such ‘assisted
killing’ strategies have been approved for clinical
trials (see review by Ockert et al. 1999).
Direct intervention to correct cancer-causing genes
is also possible. Dominantly acting genes (oncogenes)
have been targeted using antisense technology,
either with antisense transgenes, oligonucleotides
(see Carter & Lemoine 1993, Nellen & Lichtenstein
1993) or ribozymes (Welch et al. 1998, Muotri
et al. 1999). An early report of cancer gene therapy
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CHAPTER 14
with antisense oligonucleotides was that of Szczylik
et al. (1991) for the treatment of chronic myeloid
leukaemia. They used two 18-mers specific for the
BCR–ABL gene junction generated by the chromosomal translocation that causes this particular cancer,
and showed that colony formation was suppressed
in cells removed from cancer patients. Cancers
caused by loss of tumour-suppressor gene function
have been addressed by replacement strategies, in
which a functional copy of the appropriate gene is
delivered to affected cells (e.g. see Cai et al. 1993,
Harper et al. 1993, Smith et al. 1993, Hahn et al.
1996). A further strategy, known as prodrug activation therapy, involves the activation of a particular
enzyme specifically in cancer cells, which converts a
non-toxic ‘prodrug’ into a toxic product, so killing
the cancer cells. This can be achieved by driving the
expression of a so-called ‘suicide gene’ selectively
in cancer cells. An example is the HSV thymidine
kinase gene, in combination with the prodrug
ganciclovir. Thymidine kinase converts ganciclovir
into a nucleoside analogue, which is incorporated
into DNA and blocks replication by inhibiting the
DNA polymerase. Activation of the enzyme specifically in cancer cells can be achieved by preferential
delivery to dividing cells through the use of oncoretroviruses (e.g. Moolten 1986, Culver et al. 1992,
Klatzmann et al. 1996). Another way is to use
transcriptional regulatory elements that are active
only in cancer cells (e.g. Harris et al. 1994, Su et al.
1996).
The importance of SNPs
As noted earlier (p. 277), SNPs are single-base variations that occur about once every 1000 bases
along the human genome. With the availability of
the human genome sequence, a high-resolution
SNP map is being developed and this will facilitate
disease therapy in a number of ways (Rothberg
2001). Two examples will be cited here: understanding polygenic disorders and pharmacogenomics.
The disease studies cited above have all focused on
single-gene disorders. However, many of the common diseases of humans are polygenic in origin and
attempts to map genes for these complex conditions
have generally failed. The availability of SNP maps is
providing a new tool for use in genetic association
studies to identify genes for polygenic disorders, and
success has already been achieved with hypertension (Geller et al. 2000), non-insulin-dependent diabetes (Deeb et al. 2000) and cardiovascular disease
(Mason et al. 1999).
Pharmacogenomics is the term used to describe
the identification and elucidation of genetic variations that will have an impact on the efficacy of
drugs. SNPs are essential to pharmacogenomics
(McCarthy & Hilfiker 2000), for they provide an easy
means of determining the genotype of the patient.
For example, the 2-adrenergic receptor agonists
are the most widely used agents in the treatment
of asthma and several polymorphisms have been
described within the target genes. Several studies
have shown associations between SNPs in these
genes and response to therapy. One study (Buscher
et al. 1999) found that homozygotes for one allele
were up to 5.3 times more likely to respond to
albuterol than homozygotes for the other allele.
Another striking example is the response of
Alzheimer’s patients to the drug tacrine. Approximately 80% of patients not carrying the ApoE4
allele improved after tacrine treatment, whereas
60% of patients with the allele deteriorated after
treatment (Poirier et al. 1995). One scenario for the
future is that, before prescribing a drug, patients will
be genotyped by SNP analysis to determine the most
effective therapy.
Another use for pharmacogenomics is the prevention of adverse drug reactions resulting from
drug metabolism. For example, between 3 and 10%
of the Caucasian population fail to metabolize the
adrenergic-blocking drug debrisoquine and treatment
results in severe hypotension. In Afro-Americans
the frequency of this ‘poor metabolizer’ condition is
5% and in Asians it is only 1%. Affected individuals
are homozygous for a mutant cytochrome P450
gene (CYP2D6) and they also fail to metabolize over
20% of all commonly prescribed drugs, including
codeine (Gonzalez et al. 1988). The same gene also
has alleles that cause an elevated-metabolizer phenotype, which has been correlated with increased
susceptibility to cancer. Clearly, before selecting
patients for clinical trials of new drugs, it would
make sense to screen candidates for their drugmetabolizing phenotype and most large pharmaceutical companies now do this.
POGC14 9/11/2001 11:11 AM Page 293
Applications of recombinant DNA technology
Theme 3: Combating infectious disease
Introduction to theme 3
The usual way of treating bacterial and infectious
diseases is with antibiotics. As is well known, certain
microbes quickly develop resistance to the antibiotics in current use and this means that new antibiotics are required. The traditional way of obtaining
new antibiotics is the screening of new microbial
isolates from nature. An alternative way will be
described in theme 5: combinatorial biosynthesis
(p. 306). Another way is to identify new cellular
targets and screen chemical libraries for inhibitory
activities. A number of methods for identifying key
genes involved in pathogenesis have been developed
and these are described later in this section. In contrast to bacteria and fungi, viruses are not sensitive
to antibiotics and few therapies have been available.
However, this could change with the development of
antisense drugs, as described in the previous section
(p. 291). Where suitable therapies do exist, it can be
advantageous to know the identity of the pathogen
as soon as possible. Conventional laboratory procedures take several days, but PCR methodology offers a
more rapid identification, as was described on p. 274.
For many pathogens, prevention is much better than
cure, and hence vaccines are of great value. Genemanipulation techniques have greatly facilitated the
development of new vaccines, as described below.
293
cause. Finally, attenuated virus strains have the
potential to revert to a virulent phenotype upon
replication in the vaccinee. This occurs about once
or twice in every million people who receive live
polio vaccine. Recombinant DNA technology offers
some interesting solutions.
Given the ease with which heterologous genes
can be expressed in various prokaryotic and eukaryotic systems, it is not difficult to produce large quantities of purified immunogenic material for use as a
subunit vaccine. A whole series of immunologically
pertinent genes have been cloned and expressed but,
in general, the results have been disappointing. For
example, of all the polypeptides of foot-and-mouth
disease virus, only VP1 has been shown to have
immunizing activity. However, polypeptide VP1
produced by recombinant means was an extremely
poor immunogen (Kleid et al. 1981). Perhaps it is not
too surprising that subunit vaccines produced in
this way do not generate the desired immune
response, for they lack authenticity. The hepatitis B
vaccine, which is commercially available (Valenzuela
et al. 1982), differs in this respect, for expression of
the surface antigen in yeast results in the formation
of virus-like particles. A similar phenomenon is seen
with a yeast Ty vector carrying a gene for HIV coat
protein (Adams et al. 1987). These subunit vaccines
also have another disadvantage. Being inert, they
do not multiply in the vaccinee and so they do not
generate the effective cellular immune response
essential for the recovery from infectious disease.
Novel routes to vaccines
An effective vaccine generates humoral and/or cellmediated immunity, which prevents the development of disease upon exposure to the corresponding
pathogen. This is accomplished by presenting pertinent antigenic determinants to the immune system in
a fashion which mimics that in natural infections.
Conventional viral vaccines consist of inactivated,
virulent strains or live, attenuated strains, but they
are not without their problems. For example, many
viruses have not been adapted to grow to high titre
in tissue culture, e.g. hepatitis B virus. There is a
danger of vaccine-related disease when using inactivated virus, since replication-competent virus may
remain in the inoculum. Outbreaks of foot-andmouth disease in Europe have been attributed to this
Recombinant bacterial vaccines
An alternative approach to the development of
live vaccines is to start with the food-poisoning
organism Salmonella typhimurium. This organism
can be attenuated by the introduction of lesions
in the aro genes, which encode enzymes involved
in the biosynthesis of aromatic amino acids, paminobenzoic acid and enterochelins. Whereas
doses of 104 wild-type S. typhimurium reproducibly
kill mice, aro mutants do not kill mice when fed orally,
even when doses as high as 1010 organisms are
used. However, the mutant strains can establish selflimiting infections in the mice and can be detected
in low numbers in organs such as the liver and
spleen. Such attenuated strains of S. typhimurium
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CHAPTER 14
are particularly attractive as carriers of heterologous
antigens because they can be delivered orally and
because they can stimulate humoral, secretory and
cellular immune responses in the host (Charles &
Dougan 1990). Already a wide range of heterologous antigens have been expressed in such vaccine
strains (Hackett 1993).
The use of the BCG vaccine strain as an alternative vector has many advantages, including its
known safety, low cost, widespread use as a childhood vaccine and ability of a single dose to induce
long-lasting protection. Several recombinant BCG
strains have been constructed that stably express
foreign genes (Stover et al. 1991) and preliminary
results from animal studies are very encouraging.
An alternative vector is the human oral commensal
Streptococcus gordonii (Fischetti et al. 1993).
A different procedure for attenuating a bacterial
pathogen to be used as a vaccine has been proposed
by Kaper et al. (1984). They attenuated a pathogenic
strain of Vibrio cholera by deletion of DNA sequences
encoding the A1 subunit of the cholera enterotoxin.
A restriction-endonuclease fragment encoding the
A1, but not the A2 or B, sequence was deleted in vitro
from cloned cholera-toxin genes. The mutation
was then recombined into the chromosome of a
pathogenic strain. The resulting strain produces the
immunogenic but non-toxic B subunit of cholera
toxin but is incapable of producing the A subunit.
This strain has been found to be safe and immunogenic in carefully controlled clinical trials in a
number of countries (Cryz 1992).
Yet another approach to vaccine preparation has
been developed by Pizza et al. (2000), based on
whole-genome sequencing. They wanted to develop
a vaccine against group B meningococci but were
hampered by the fact that there was considerable
sequence variation in surface-exposed proteins.
Starting with the entire genome sequence of a group
B strain of Neisseria meningitidis, they identified 350
proteins as potential protective antigens. All 350
candidate antigens were expressed in E. coli, purified
and used to immunize mice. The sera from the mice
allowed the identification of proteins that are surfaceexposed in meningococci, that are conserved across
a range of strains and that induce a bactericidal antibody response, a property known to correlate with
vaccine efficacy in humans.
Recombinant viruses as vaccines
Recombinant viruses can be used as vectors to
express heterologous antigens, and thus function as
live vaccines. The first animal virus to be exploited in
this way was vaccinia, which had been used previously as a non-recombinant vaccine providing
cross-protection against variola virus, the causative
agent of smallpox. Contributing to its success as a
live vaccine were its stability as a freeze-dried preparation, its low production cost and the ability to
administer the vaccine by simple dermal abrasion.
Vaccinia remains the most widely explored recombinant viral vaccine, and many antigens have been
expressed using this vector, essentially using the
methods described on p. 197 (Ulaeto & Hruby 1994,
Moss 1996). The most successful recombinant-virus
vaccination campaign to date involved the use of
recombinant vaccinia virus expressing rabies-virus
glycoprotein. This was administered to the wild population of foxes in central Europe by providing a bait
consisting of chicken heads spiked with the virus.
The epidemiological effects of vaccination were most
evident in eastern Switzerland, where two decades
of rabies came to a sudden end after only three vaccination campaigns (Brochier et al. 1991, Flamand
et al. 1992), and in Belgium, where the disease was
all but eliminated (Brochier et al. 1995).
One disadvantage of this approach in humans is
the unacceptably high risk of adverse reactions to
the vaccine. For use in humans, vaccinia must be further attenuated to make it replication-deficient and
to minimize the likelihood of replication-competent
viruses arising by recombination (e.g. Cooney et al.
1991, Sutter & Moss 1992, Tartaglia et al. 1992).
Highly attenuated vaccinia derivatives (Table 14.6)
have been used to express a range of viral, bacterial
and parasite antigens, with some reaching clinical
trials (reviewed by Moss 1996, Paoletti 1996). Other
poxviruses, such as canarypox, have also been
developed as potential vaccines (e.g. see Perkus et al.
1995, Fries et al. 1996, Myagkikh et al. 1996).
As well as vaccinia, adenovirus and alphavirus
vectors have also been investigated as potential
recombinant vaccines. Adenovirus has been used to
express various antigens, including HSV and rabies
virus glycoproteins (Gallichan et al. 1993, Xiang
et al. 1996). Among the alphaviruses, Semliki Forest
POGC14 9/11/2001 11:11 AM Page 295
Applications of recombinant DNA technology
295
Table 14.6 Immune response to heterologous antigens expressed by vaccinia virus recombinants.
Antigen
Neutralizing antibodies
Cellular immunity
Animal protection
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
not determined
Rabies virus glycoprotein
Vesicular stomatitis glycoprotein
Herpes simplex virus glycoprotein D
Hepatitis B surface antigen
Influenza virus haemagglutinin
Human immunodeficiency virus envelope
Reproduced with permission from Tartaglia & Paoletti (1988).
Table 14.7 A selection of recombinant
vaccines against animal viruses
produced in plants.
Antigen
Herpes virus B surface antigen
Rabies glycoprotein
Norwark virus coat protein
Foot-and-mouth virus VP1
Cholera toxin B subunit
Human cytomegalovirus
glycoprotein B
virus has been used to express the gp160 protein
of HIV (Berglund et al. 1997) and Sindbis virus
has been used to express antigens from Japanese
encephalitis virus (Pugachev et al. 1995).
Recombinant proteins expressed by viruses have
been shown to be immunogenetic. However, stronger
stimulation of the immune response can often be
achieved by presenting the antigen on the surface of
the virus attached to a host virus-derived carrier
protein. The advantage of this strategy is that the
recombinant antigen is presented as multiple copies.
This strategy is borrowed from the surface display
of foreign antigens on bacteria that are used as live
vaccines (see above) but is much safer. Indeed, while
a number of conventional viral vaccines, such as
vaccinia, have been developed as surface-display
systems (e.g. see Katz & Moss 1997, Katz et al. 1997),
even plant viruses can be used for this purpose,
as discussed below (Porta & Lomonossoff 1996,
Johnson et al. 1997).
Host-plant system
Reference
Tobacco
Tomato
Tobacco, potato
Arabidopsis
Potato
Tobacco
Mason et al. 1992
McGarvey et al. 1995
Mason et al. 1996
Carrillo et al. 1998
Arakawa et al. 1998
Tackaberry et al. 1999
Plants as edible vaccines
Plants have been explored as a cheap, safe and
efficient production system for subunit vaccines
(Table 14.7), with the added advantage that orally
administered vaccines can be ingested by eating the
plant, therely eliminating the need for processing
and purification (reviewed by Mason & Arntzen
1995, Walmsey & Arntzen 2000). The earliest
demonstration was the expression of a surface antigen from the bacterium S. mutans in tobacco. This
bacterium is the causative agent of dental caries,
and it was envisaged that stimulation of a mucosal
immune response would prevent the bacteria colonizing the teeth and therefore protect against tooth
decay (Curtis & Cardineau, 1990).
A number of edible transgenic plants have been
generated expressing antigens derived from animal
viruses. For example, rabies glycoprotein has been
expressed in tomato (McGarvey et al. 1995), hepatitis
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CHAPTER 14
B virus antigen in lettuce (Ehsani et al. 1997) and
cholera antigen in potato (Arakawa et al. 1997). As
well as animal virus antigens, autoantigens associated with diabetes have also been produced (Ma et al.
1997, Porceddu et al. 1999). Plants have also
been infected with recombinant viruses expressing
various antigen epitopes on their surfaces. Cowpea
mosaic virus (CMV) has been extensively developed
as a heterologous antigen-presenting system (Porta
et al. 1994, Lomonossoff & Hamilton 1999). There
have been some recent successes in vaccination
trials using recombinant CMV vectors expressing
epitopes of HIV gp41 (McLain et al. 1995, 1996a,b)
and canine parvovirus (Dalsgaard et al. 1997). The
first clinical trials using a plant-derived vaccine
were conducted in 1997 and involved the ingestion
of transgenic potatoes expressing the B subunit of
the E. coli heat-labile toxin, which causes diarrhoea.
This resulted in a successful elicitation of mucosal
immunity in test subjects (Tacket et al. 1994).
DNA vaccines
The immune system generates antibodies in response
to the recognition of proteins and other large molecules carried by pathogens. In each of the examples
above, the functional component of the vaccine
introduced into the host is the protein responsible
for the elicitation of the immune response. The introduction of DNA into animals does not generate an
immune response against the DNA molecule, but, if
that DNA is expressed to yield a protein, that protein
can stimulate the immune system. This is the basis
of DNA vaccination, as first demonstrated by Ulmer
et al. (1993). DNA vaccines generally comprise a
bacterial plasmid carrying a gene encoding the
appropriate antigen under the control of a strong
promoter that is recognized by the host cell. The
advantages of this method include its simplicity, its
wide applicability and the ease with which large
quantities of the vaccine can be produced. The DNA
may be administered by injection, using liposomes
or by particle bombardment. In the original demonstration, Ulmer and colleagues introduced DNA
corresponding to the influenzavirus nucleoprotein
and achieved protection against influenza infection.
Since then, many DNA vaccines have been used
to target viruses (e.g. measles (Cardoso et al. 1996);
HIV (Wang et al. 1993, Fuller et al. 1997, Hinkula
et al. 1997); Ebola virus (Xu et al. 1998) ), other
pathogens (e.g. tuberculosis (Huygen et al. 1996) )
and even the human cellular prion protein in mice
(Krasemann et al. 1996).
The DNA-vaccination approach has several additional advantages. These include the following:
• Certain bacterial DNA sequences have the innate
ability to stimulate the immune system (see Klinman
et al. 1997, Roman et al. 1997).
• Other genes encoding proteins influencing the function of the immune response can be co-introduced
along with the vaccine (e.g. Kim et al. 1997).
• DNA vaccination can be used to treat diseases that
are already established as a chronic infection (e.g.
Mancini et al. 1996).
In principle, DNA vaccination has much in
common with gene therapy (discussed above), since
both processes involve DNA transfer to humans,
using a similar selection of methods. However, while
the aim of gene therapy is to alleviate disease, by
either replacing a lost gene or blocking the expression of a dominantly acting gene, the aim of DNA
vaccination is to prevent disease, by causing the
expression of an antigen that stimulates the immune
system.
Selecting targets for new
antimicrobial agents
In attempting to develop new antimicrobial agents,
including ones that are active against intractable
pathogens such as the malarial parasite, it would be
useful to know which genes are both essential for
virulence and unique to the pathogen. Once these
genes have been identified, chemical libraries can be
screened for molecules that are active against the
gene product. Two features of this approach deserve
further comment. First, inhibition of the gene product could attenuate the organism’s virulence but
would not result in death of the organism in vitro;
that is, no effect would be seen in whole-organism
inhibition assays and so molecules only active in
vivo would be missed. Secondly, target genes can be
selected on the basis that there are no human counterparts. Thus, active molecules will be less likely
to be toxic to humans. A number of different approaches have been developed for identifying virulence
POGC14 9/11/2001 05:28 PM Page 297
Applications of recombinant DNA technology
determinants, and details are presented below for
three of these: in vivo expression technology (IVET),
differential fluorescence induction and signaturetagged mutagenesis.
In vivo expression technology (IVET)
IVET was developed to positively select those genes
that are induced specifically in a microorganism
297
when it infects an animal or plant host (Mahan et al.
1993). The basis of the system is a plasmid carrying
a promoterless operon fusion of the lacYZ genes
fused to the purA or thyA genes downstream of a
unique BglII cloning site (Fig. 14.11). This operon
fusion was constructed in a suicide-delivery plasmid. Cloning of pathogen DNA into the BglII site
results in the construction of a pool of transcriptional fusions driven by promoters present in the
Bgl II
thyA
+
lacZ
IVET
plasmid
Salmonella DNA
Cut with Bgl II
and ligate
Pl
thyA
lacZ
Transfer to thyA strain of Salmonella.
Grow in presence of thymine
Isolate pathogen on
media + thymine + Xgal
Fig. 14.11 The basic principle of IVET.
See text for details.
White
colonies
Blue
colonies
thyA expressed
from promoter
only active in vivo
thyA expressed
from promoter
active in vitro
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CHAPTER 14
cloned DNA. The pool of fusions is transferred to
a strain of the pathogen carrying a purA or thyA
deletion and selection made for integration into the
chromosome. The recombinant pathogen is then
used to infect a test animal. Fusions that contain
a promoter that is active in the animal allow
transcription of the purA or thyA gene and hence
bacterial survival. When the surviving pathogens
are reisolated from the test animal, they are tested in
vitro for their levels of β-galactosidase. Clones that
contain fusions to genes that are specifically induced
in the test animal show little lacZ expression on
laboratory media and DNA sequencing can be used
to identify the genes to which these promoters belong.
The IVET system was initially developed for use in
a murine typhoid model. Since it was first described,
IVET has been used with a wide variety of Grampositive and Gram-negative organisms, including
‘difficult’ organisms, such as Mycobacterium. One
disadvantage of the original IVET system is that it
inherently selects for promoters that are constitutively and highly expressed in vivo. An alternative
IVET system uses tnpR operon fusions in place of
purA or thyA. The tnpR locus encodes a site-specific
resolvase and is used as the selection gene. Expression of this gene results in resolvase synthesis
and the deletion of a resolvase-specific reporter.
The advantage of this system is that it can be used
to identify those genes that are expressed only
transiently during animal infection (Camilli &
Mekalanos 1995, Camilli 1996). Handfield and
Levesque (1999) have described a number of other
modifications to the IVET technology.
Differential fluorescence induction
This technique represents a different way of identifying environmentally controlled promoters and was
originally developed to facilitate identification of
Salmonella genes that are differentially expressed
with macrophages (Valdivia & Falkow 1997). Fragments of DNA from Salmonella were fused to a promotorless gene for green fluorescent protein (GFP)
and returned to the Salmonella, which was then used
to infect macrophages. Cells that became fluorescent
were recovered using a fluorescence-activated cell
sorter (FACS) and grown on media in the absence of
macrophages. Bacteria that were non-fluorescent in
the extracellular environment were sorted and used
for a second round of macrophage infection. Bacteria
still capable of generating fluorescent macrophages
were found to contain GFP fusions that were
up-regulated by the macrophage’s intracellular
environment.
Signature-tagged mutagenesis
This technique is a variation on the use of transposon mutagenesis that has been applied to the
identification of bacterial virulence genes. The basic
principle is to create a large number of different
transposon-generated mutants of a pathogenic
organism and to identify those mutants that can
survive in vitro but not in vivo – that is, it is a wholegenome scan for habitat-specific genes. Although
one could test each mutant individually, this would
be very laborious and would use large numbers
of animals. By using signature-tagged mutagenesis,
one can test large numbers of different mutants
simultaneously in the same animal. This is achieved by
tagging each transposon mutant with a different
DNA sequence or tag.
The use of sequence-tagged mutagenesis was first
described for the identification of virulence genes
from S. typhimurium in a mouse model of typhoid
fever (Hensel et al. 1995), as shown in Fig. 14.12.
The tags comprise different sequences of 40 bp
[NK]20, where N = A, C, G or T and K = G or T. The
arms were designed so that the amplification of the
tags by PCR with specific primers would produce
probes with 10 times more label in the central region
than in each arm. The double-stranded tags were
ligated into a Tn5 transposon and transferred from
E. coli to S. typhimurium by conjugation. A library of
over 1500 exconjugants resulting from transposition events was stored in microtitre dishes. Of these
exconjugants, 1152 were selected and prepared as
12 pools of 96 mutants. Each pool was injected into
the peritoneum of a different mouse and infection
allowed to proceed. Bacteria were then recovered
from each mouse by plating spleen homogenates
on culture media. DNA was extracted from the
recovered bacteria and the tags in this DNA were
amplified by PCR. Those tags present in the initial
pool of bacteria but missing from the recovered
media represent mutations in genes essential for
POGC14 9/11/2001 11:11 AM Page 299
Applications of recombinant DNA technology
1 Construct 1017 oligonucleotide tags with
the structure
(NK)20
2 Clone tags into transposon Tn5 resident in plasmid vector.
3 Transform tagged transposon plasmids into E.coli selecting
for kanamycin resistance. The individual transformants are
not isolated but kept as a pool.
299
Environomics
The techniques described above to identify bacterial
virulence genes can be applied to other microbe–
host interactions, e.g. protozoan infections of humans,
bacterial or fungal pathogenesis of plants, or even
beneficial rhizosphere–microbe interactions (Rainey
1999). Essentially, these techniques are methods
for scanning the entire microbial genome for genes
that are expressed under particular environmental conditions. This approach has been termed
‘environomics’.
Theme 4: Protein engineering
4 Conjugate plasmids into Salmonella and select for transposition
of tagged transposon. Each kanamycin resistant cell should
carry a different tag.
5 Arrange each individual kanamycin resistant clone, which
should be result of different insertion event and will have
different tag, in 8 ×12 grid in 96 well microtitre plates.
6 Pool groups of 96 clones and infect mouse. After period for
infection to be established, re-isolate kanamycin resistant cells
from mouse. Extract DNA.
7 Screen DNA from mouse isolates and determine which tags
are absent relative to those that are present in microtitre trays.
Tags that are missing must be attached to transposons that
have inserted into a gene essential for mouse virulence.
Fig. 14.12 The basic methodology for signature tagged
mutagenesis.
virulence. In this way 28 different mutants with
attenuated virulence were identified and some of
these mutants were in genes not previously identified.
The principle of signature-tagged mutagenesis
has been extended to the analysis of pathogenicity
determinants in a wide range of bacteria (for a review,
see Handfield & Levesque 1999) and to fungi (Brookman & Denning 2000).
Introduction to theme 4
One of the most exciting aspects of recombinant
DNA technology is that it permits the design, development and isolation of proteins with improved
operating characteristics and even completely novel
proteins. The simplest example of protein engineering involves site-directed mutagenesis to alter key
residues, as originally shown by Winter and colleagues (Winter et al. 1982, Wilkinson et al. 1984).
From a detailed knowledge of the enzyme tyrosyltRNA synthetase from Bacillus stearothermophilus,
including its crystal structure, they were able to
predict point mutations in the gene that should
increase the enzyme’s affinity for the substrate ATP.
These changes were introduced and, in one case, a
single amino acid change improved the affinity for
ATP by a factor of 100. Using a similar approach, the
stability of an enzyme can be increased. Thus Perry
and Wetzel (1984) were able to increase the thermostability of T4 lysozyme by the introduction of a
disulphide bond. However, although new cysteine
residues can be introduced at will, they will not
necessarily lead to increased thermal stability (Wetzel
et al. 1988).
Improving therapeutic proteins with
single amino acid changes
As noted earlier (p. 283), many recombinant
proteins are now being used therapeutically. With
some of them, protein engineering has been used to
generate second-generation variants with improved
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pharmacokinetics, structure, stability and bioavailability (Bristow 1993). For example, in the neutral
solutions used for therapy, insulin is mostly assembled
as zinc-containing hexamers. This self-association
may limit absorption. By making single amino acid
substitutions, Brange et al. (1988) were able to
generate insulins that are essentially monomeric at
pharmaceutical concentrations. Not only have
these insulins preserved their biological activity, but
they are also absorbed two to three times faster.
Similarly, replacing an asparagine residue with
glutamine altered the glycosylation pattern of TPA.
This in turn significantly increased the circulatory
half-life, which in the native enzyme is only 5 min
(Lau et al. 1987). Proteins can also be engineered to
be resistant to oxidative stress, as has been shown
with AAT (see Box 14.3).
Improving enzymes: subtilisin as a
paradigm for protein engineering
Proof of the power of gene manipulation coupled
with the techniques of in vitro (random and sitedirected) mutagenesis as a means of generating
improved enzymes is provided by the work done on
subtilisin over the past 15 years (for review, see
Bryan 2000). Every property of this serine protease
has been altered, including its rate of catalysis, substrate specificity, pH-rate profile and stability to
oxidative, thermal and alkaline inactivation. In the
process, well over 50% of the 275 amino acids of
subtilisin have been changed. At some positions in
the molecule, the effects of replacing the usual
amino acid with all the other 19 natural amino acids
have been evaluated.
Box 14.3 Oxidation-resistant variants of a1-antitrypsin (AAT)
Cumulative damage to lung tissue is thought to be
responsible for the development of emphysema, an
irreversible disease characterized by loss of lung
elasticity. The primary defence against elastase
damage is AAT, a glycosylated serum protein of
394 amino acids. The function of AAT is known
because its genetic deficiency leads to a premature
breakdown of connective tissue. In healthy
individuals there is an association between AAT and
neutrophil elastase followed by cleavage of AAT
between methionine residue 358 and serine
residue 359 (see Fig. B14.2).
After cleavage, there is negligible dissociation of
the complex. Smokers are more prone to emphysema,
because smoking results in an increased
concentration of leucocytes in the lung and
Met 358
394
consequently increased exposure to neutrophil
elastase. In addition, leucocytes liberate oxygen
free radicals and these can oxidize methionine358 to methionine sulphoxide. Since methionine
sulphoxide is much bulkier than methionine, it does
not fit into the active site of elastase. Hence oxidized
AAT is a poor inhibitor. By means of site-directed
mutagenesis, an oxidation-resistant mutant of AAT
has been constructed by replacing methionine-358
with valine (Courtney et al. 1985). In a laboratory
model of inflammation, the modified AAT was an
effective inhibitor of elastase and was not inactivated
by oxidation. Clinically, this could be important,
since intravenous replacement therapy with plasma
concentrates of AAT is already being tested on
patients with a genetic deficiency in AAT production.
Ser 359
394
Met 358
Fig. B14.2 The cleavage of α1-antitrypsin on
binding to neutrophil elastase.
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Applications of recombinant DNA technology
Many of the changes described above were made
to improve the ability of subtilisin to hydrolyse protein when incorporated into detergents. However,
serine proteases can be used to synthesize peptides
and this approach has a number of advantages over
conventional methods (Abrahmsen et al. 1991).
A problem with the use of subtilisin for peptide
synthesis is that hydrolysis is strongly favoured over
aminolysis, unless the reaction is undertaken in
organic solvents. Solvents, in turn, reduce the halflife of subtilisin. Using site-directed mutagenesis, a
number of variants of subtilisin have been isolated
with greatly enhanced solvent stability (Wong et al.
1990, Zhong et al. 1991). Changes introduced
included the minimization of surface changes to reduce
solvation energy, the enhancement of internal polar
and hydrophobic interactions and the introduction
of conformational restrictions to reduce the tendency
of the protein to denature. Designing these changes
requires an extensive knowledge of the enzyme’s
structure and function. Chen and Arnold (1991,
1993) have provided an alternative solution. They
utilized random mutagenesis combined with screening for enhanced proteolysis in the presence of
solvent (dimethyl formamide) and substrate (casein).
The engineering of subtilisin has now gone one
step further, in that it has been modified such that
aminolysis (synthesis) is favoured over hydrolysis,
even in aqueous solvents. This was achieved by
changing a serine residue in the active site to cysteine (Abrahmsen et al. 1991). The reasons for this
enhancement derive mainly from the increased
affinity and reactivity of the acyl intermediate for the
amino nucleophile (Fig. 14.13). These engineered
‘peptide ligases’ are in turn being used to synthesize
novel glycopeptides. A glycosyl amino acid is used
in peptide synthesis to form a glycosyl peptide ester,
H2O
RCOOH + E
OH
Hydrolysis
RCOOR’ + E
OH
RCOO E
R–NH2
Aminolysis
RCOOR’ + E
Fig. 14.13 The aminolysis (synthetic) and hydrolysis
reactions mediated by an acylated protease.
OH
301
which will react with another C-protected peptide
in the presence of the peptide ligase to form a larger
glycosyl peptide.
Methods for engineering proteins:
the rational approach
Given the range of modifications made to subtilisin,
the question is not ‘what modifications are possible?’
(for review, see Arnold 2001) but ‘how do I achieve
the modifications that I wish to make?’ There are
two general techniques: the rational approach and
‘directed evolution’. There are two variations on the
rational approach. The first of these makes use of
comparisons between related proteins. For example,
barnase and binase are ribonucleases that have 85%
sequence identity but differ in their thermal stability.
Using site-directed mutagenesis, Serrano et al. (1993)
tested the impact on thermostability of barnase of
all 17 amino acid differences between barnase and
binase. They observed effects between +1.1 and −1.1
kcal/mol and a multiple mutant combining six of
the substitutions displayed a stability increase of
3.3 kcal/mol over the wild-type enzyme.
The second variation on the rational approach
requires that the three-dimensional structure of the
protein be known. Based on an analysis of this structure, particular residues are selected for mutagenesis
and the biological effects then evaluated. That this
approach works was clearly shown by the early
work on lysozyme and subtilisin, but the chances of
success are not predictable. One disadvantage of this
approach is illustrated by the results of Spiller et al.
(1999), who were investigating the thermostability
of an esterase using directed evolution. When they
evaluated the effect of certain mutations, they
concluded that no amount of rational analysis of
the crystal structure would have led them to predict
the results obtained.
Protein engineering through
directed evolution
Directed evolution involves repeated rounds of
random mutagenesis, followed by selection for the
improved property of interest. Any number of
methods can be used to introduce the mutations,
including mutagenic base analogues, chemical
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mutagenesis, error-prone PCR and spiked synthetic
oligonucleotides. The key element in the process is
the ability to screen large numbers of mutants. An
example is the isolation of a more thermostable
subtilisin. Up to 1000 mutant clones are gridded
out on replica plates and, once they are grown, one
plate is incubated at an elevated temperature long
enough to inactivate the wild-type enzyme. When
an assay for hydrolytic activity is subsequently
performed, only mutants with stability greater than
that of the wild type will display measurable activity.
Once stable mutants have been identified, the replicate colony can be grown to identify the mutation.
Once stabilizing single amino acid changes have
been identified, building a highly stable subtilisin
can be accomplished by combining individual amino
acids into the same molecule (Pantoliano et al. 1988,
1989, Zhao & Arnold 1999).
One of the disadvantages of the screening method
described above is that it is labour-intensive and the
maximum feasible number of mutants that can be
examined in a single screen is 104–105. By combin-
ing mutagenesis with phage display, up to 109 different mutants can theoretically be screened in a single
experiment. The constraint with this approach is
that phage display is really best suited to selecting
proteins with altered binding characteristics, rather
than the other properties one might wish to engineer. Despite this, a number of groups (Atwell &
Wells 1999, Demartis et al. 1999, Olsen et al. 2000)
were able to select variant proteases with novel
substrate specificities.
Gene families as aids to
protein engineering
An alternative approach to directed evolution is
‘DNA shuffling’, which is also known as ‘molecular
breeding’ (Minshull & Stemmer 1999, Ness et al.
2000). This method can only be adopted if the target
protein belongs to a known protein family. If it does,
the genes for the different family members are
isolated and artificial hybrids created (Fig. 14.14).
As an example of this approach, Ness et al. (1999)
Ancestral species
Evolution
Species 1
Species 2
Species 3
Species 4
DNA shuffling
in vitro
Hybrid genes
Fig. 14.14 Schematic representation of
gene shuffling.
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Applications of recombinant DNA technology
started with the genes for 26 members of the subtilisin family and created a library of chimeric
proteases. When this library was screened for four
distinct enzyme properties, variants were found that
were significantly improved over any of the parental
enzymes for each individual property. In a similar
way, Powell et al. (2000) were able to select
retroviruses with increased stability for use as
gene-therapy vectors.
As a final twist on the methods for engineering
proteins, Lehmann et al. (2000) have combined the
use of protein families with a rational approach.
They started with a family of mesophilic phytases
whose amino acid sequences had been determined.
Using these data they constructed a ‘consensus’
phytase sequence and found that enzyme with this
sequence was much more thermostable than any of
the parent enzymes. Refinement of the approach
combined with site-directed mutagenesis resulted in
the isolation of an enzyme whose unfolding temperature was 27–34°C higher than that of the parent
enzymes. Whereas all the methods described earlier
tend to yield mutants with single amino acid changes,
this method generated variants with multiple amino
acid changes spread throughout the molecule.
Theme 5: Metabolic engineering
Introduction to theme 5
When the large-scale production of penicillin was
begun in the 1940s, the yields of penicillin were
measured in micrograms per litre of culture. Demand
for the antibiotic was outstripping supply and higheryielding strains were badly needed. Since nothing
was known about the biosynthetic pathway, a programme of strain improvement was set in place that
involved random mutation and screening. The best
strain from each cycle of improvement then became
the starting-point for the next round of selection. In
this way the yield of penicillin was steadily increased
until it reached the tens of grams per litre that can be
achieved today. As each new antibiotic was discovered, the same process of strain improvement was
applied. In every case, the biochemical and genetic
basis of the beneficial mutations was not known.
Only when the details of gene regulation and
metabolic-pathway regulation (allosteric control)
303
had been elucidated could we even begin to understand how antibiotic yields might have been improved.
Based on our current knowledge of metabolic
regulation we can predict that the changes in the
improved strains described above will involve all of
the following:
• Removal of rate-limiting transcriptional and allosteric controls.
• Kinetic enhancement of rate-limiting enzymes.
• Genetic blockage of competing pathways.
• Enhanced carbon commitment to the primary
metabolic pathway from central metabolism.
• Modification of secondary metabolic pathways to
enhance energy metabolism and availability of
enzymatic cofactors.
• Enhanced transport of the compound out of the cell.
Today, if one starts with a wild-type strain and
wants to turn it into an overproducer, then the
approach would be to use recombinant DNA technology to make these desired changes in a rational
way, as exemplified below by phenylalanine. There
are many other examples of rational ‘metabolic
engineering’ and these have been reviewed by
Chotani et al. (2000).
Designed overproduction of
phenylalanine
Phenylalanine is a key raw material for the synthesis
of the artificial sweetener aspartame. Phenylalanine
can be synthesized chemically but is too expensive if
made this way. In the 1980s bacterial strains that
overproduced phenylalanine were developed, using
the traditional mutation and selection method. At
the same time, a programme of rational strain development was instituted at G.D. Searle, the company
who owned the patent for aspartame. The startingpoint for this programme was an analysis of the
biosynthetic pathway (Fig. 14.15). Removing feedback inhibition of key steps is an essential first
step. In the case of a phenylalanine producer, it is
essential to knock out any feedback inhibition of the
pathway from chorismate to phenylalanine and this
was achieved by selecting strains resistant to phenylalanine analogues. The conversion of erythrose-4phosphate (E4P) and phosphoenol pyruvate (PEP)
to DAHP is also subject to feedback inhibition, but,
since there are three different enzymes here, each
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Erythrose-4-phosphate (E4P) + phosphoenol pyruvate (PEP)
DPAH
Chorismate
Prephenate
Anthranilate
Indoleglycerol phosphate
Tyrosine
Phenylalanine
Repression
Feedback inhibition
inhibited by a different aromatic end-product, all
that is necessary for a phenylalanine overproducer
is to clone the tryptophan-sensitive enzyme and
have it overexpressed. To overcome repression of
enzyme synthesis, the existing promoters were
removed and replaced with one that could be controlled more easily in industrial-scale fermentations.
The above changes removed the natural control
circuits. The next step was to remove competing
pathways, i.e. the synthesis of tyrosine and tryptophan. This was easily achieved by making a tyrosine
and tryptophan double auxotroph. Note that stable
(non-reverting) auxotrophs can best be made by
deleting part or all of the relevant genes. This is a
task that is easy using recombinant DNA technology. Once all the control circuits and competing
pathways had been removed, attempts were made to
increase the carbon flux through the biosynthetic
pathway. Surprisingly, overexpressing all the genes
in the pathway did not enhance the yield of phenylalanine. One explanation was that the supply of precursors (E4P and/or PEP) was rate-limiting. This
Tryptophan
Fig. 14.15 The regulation of the
biosynthesis of aromatic amino acids. Note
that indoleglycerol phosphate is converted
to tryptophan via indole but that the indole
normally is not released from the tryptophan
synthase complex.
was confirmed when cloning transketolase (to
enhance E4P levels) and eliminating pyruvate kinase
(to enhance PEP levels) enhanced yields.
New routes to small molecules
Recombinant DNA technology can be used to
develop novel routes to small molecules. Good
examples are the microbial synthesis of the blue dye
indigo (Ensley et al. 1983) and the black pigment
melanin (Della-Cioppa et al. 1990). Neither compound is produced in bacteria. The cloning of a
single gene from Pseudomonas putida – that encoding
naphthalene dioxygenase – resulted in the generation of an E. coli strain able to synthesize indigo in a
medium containing tryptophan (Fig. 14.16). Similarly, cloning a tyrosinase gene in E. coli led to
conversion of tyrosine to dopaquinone, which spontaneously converts to melanin in the presence of air.
To overproduce these compounds, one generates a
strain of E. coli that overproduces either tryptophan
or tyrosine, rather than phenylalanine, as described
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Applications of recombinant DNA technology
305
NH2
CH2
CH
COOH
N
Tryptophan
Tryptophanase (E.coli )
N
Indole
Naphthalene dioxygenase
(cloned)
H
OH
H
OH
N
cis-indole-2, 3-dihydrodiol
O
N
O
NH2
Isatin
Air oxidation
Isatic acid
Air oxidation
O
O
N
N
Fig. 14.16 The biosynthesis of indigo
in E. coli and the formation of
alternative end products.
OH
O
N
Indoxyl
O
Isatin
hydrolase
OH
Air
oxidation
Spontaneous
N
O
Indigo
above. With both indigo and melanin, yields are
improved by increasing the levels of cofactors. Also,
in the case of indigo biosynthesis, it is necessary to
engineer the tryptophan synthase gene. The reason
for this is that indole is an intermediate in the
biosynthesis of tryptophan (Fig. 14.15). However,
normally it is not free in the cytoplasm but remains
trapped within the tryptophan synthase complex.
By modifying the trpB gene, encoding the subunit of
O
N
Indirubin
tryptophan synthase, it was possible for the indole
to be released for conversion by the dioxygenase
(Murdock et al. 1993).
One disadvantage of the new route to indigo is
that one of the intermediates in its synthesis,
indoxyl, can undergo an alternative spontaneous
oxidation to isatin and indirubin. The latter compound is an isomer of indigo with similar dyeing
properties, but instead of being blue it is a deep
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(a)
Chemical
D-glucose
Chemical
Ascorbic acid
D-sorbitol
2 keto-Lgulonic acid
Acetobacter
suboxydans
Chemical
L-sorbose
Diacetone
-2 keto-Lgulonic acid
(b)
CHO
COOH
OH
O
OH
Erwinia
enzymes
OH
CH2OH
D-glucose
HO
O
2,5 DKG
reductase
Endogenous
HO
COOH
(Corynebacterium)
HO
OH
O
CH2OH
2,5 diketo-Dgluconic acid
burgundy colour. To make textile-quality indigo,
there must be no indirubin present. Screening soil
microorganisms with the capacity to degrade indole
resulted in the identification of an enzyme, isatin
hydrolase, that can degrade isatin to isatic acid.
After cloning the gene for isatin hydrolase in the
indigo overproducing strains, the indigo product
obtained performed as well as chemically produced
material.
A slightly different approach to that above has
yielded a new route to vitamin C. The conventional
process starts with glucose and comprises one
mirobiological and four chemical steps (Fig. 14.17).
By cloning in Erwinia a single gene – that from
Corynebacterium encoding 2,5-diketogluconic acid
reductase – the process can be simplified to a single
microbiological and a single chemical step (Anderson et al. 1985). After observations of unexpectedly
low yields of 2-ketogulonic acid in the recombinant
OH
HO
CH2OH
2-keto-Lgulonic acid
Fig. 14.17 Simplified route to vitamin
C (ascorbic acid) developed by cloning
in Erwinia the Corynebacterium gene for
2,5-diketogluconic acid reductase.
(a) Classical route to vitamin C. (b) The
simplified route to 2-ketogulonic acid,
the immediate precursor of vitamin C.
strain, it was found that 2-ketogulonic acid was
converted to l-idonic acid by an endogenous 2ketoaldonate reductase. Cloning, deletion mutagenesis and homologous recombination of the mutated
reductase gene into the chromosome were some
of the several steps taken to develop an organism
capable of accumulating large amounts (120 g/l) of
2-ketogulonic acid (Lazarus et al. 1990). So far,
attempts to manufacture vitamin C directly from
glucose have been unsuccessful. However, enzymes
that can convert 2-ketogulonic acid to ascorbic acid
have been identified and the objective now is to
clone these activities into Erwinia (Chotani et al.
2000).
Combinatorial biosynthesis
A number of widely used antibiotics and immunosuppressants belong to a class of molecules known
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Applications of recombinant DNA technology
307
O
H3C
CH3
OH
H3C
O
OH
O
OH
CH3
OH
OH
Sugar
H3C
O
CH3
CH3
OCH3 O
OH
Sugar
Sugar
O
CH3
Erythromycin
Daunorubicin
OCH3
HO
OCH3
O
H3C
O
CH3
H
O
H3C
CH3
H
O
O
H
H
O
H3C
O
O
OH
H
CH3
H
CH3
O
CH3
H
OCH3
Avermectin
O
CH3
O
HC
H3C
H3C
O
Sugar
O
Sugar
OH
CH3
Fig. 14.18 Some examples of
polyketides.
Tylosin
as polyketides. These molecules, which are synthesized by actinomycetes, have a fairly complex
structure (Fig. 14.18). The genes involved in the
biosynthesis of polyketides are clustered, thereby
facilitating the cloning of all of the genes controlling
the synthetic pathway. The first cluster (the act genes)
to be cloned was that for actinorhodin. When parts
of the act gene cluster were introduced into streptomycetes making related polyketides, completely new
antibiotics were produced (Hopwood et al. 1985).
For example, introducing the actVA gene from Streptomyces coelicolor into a strain that makes medermycin
leads to the synthesis of mederrhodinA (Fig. 14.19).
This approach has been repeated many times since
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H
O
Actinorhodin
O
previous theme (p. 302), and this approach has
been widely adopted (Baltz 1998). However, novel
polyketides can also be generated by simply changing the order of the different activities in type I
synthases (McDaniel et al. 1999).
CH3
O
COOH
O
H
Medermycin
Engineering metabolic control over
recombinant pathways
N(CH2)2
H
O
HO
H3C
O
O
CH3
O
O
H
O
O
O
actVA gene
Mederrhodin
N(CH2)2
H
O
HO
H3C
O
CH3
O
O
O
H
O
O
O
Fig. 14.19 The formation of the new antibiotic mederrhodin
from medermycin by the actVA gene product.
with other polyketides (for review, see Baltz 1998)
and is known as combinatorial biosynthesis.
Once a number of polyketide biosynthetic gene
clusters had been cloned and sequenced, new
insights were gained on the mechanism of synthesis.
In particular, two enzymic modes of synthesis were
discovered. In particular, polyketide synthesis takes
place on an enzyme complex in a manner analogous
to fatty acid synthesis. Furthermore, there are two
types of complex. In type II complexes, the different
enzymic activities are encoded by separate subunits.
In contrast, in type I synthesis all the different
enzyme activities are encoded by a single, very large
gene. Clearly, the polyketide synthases are prime
candidates for DNA shuffling, as described in the
When a recombinant cell overproduces a protein or
the components of a biosynthetic pathway, there is a
marked reduction in metabolic activity, coupled with
a retardation of growth (Kurland & Dong 1996).
These phenotypic characteristics result from the fact
that the constant demands of production placed
upon the cell interfere with its changing requirements for growth. To alleviate this problem, Farmer
and Liao (2000) designed a dynamic control circuit
that is able to sense the metabolic state of the cell and
thereby regulate the expression of a recombinant
pathway. This approach is termed ‘metabolic control
engineering’.
An essential component of a dynamic controller is
a signal that will reflect the metabolic state of the
cell. Acetyl phosphate was selected as the signal,
since it is known to be a regulator of various operons
influenced by nutrient availability. A component of
the Ntr regulon in E. coli, NRI, is capable of sensing
the acetyl phosphate level in the cell. When phosphorylated by acetyl phosphate, it is capable of
binding to the glnAp2 promoter and activating
transcription (Fig. 14.20). To reconstruct this
control module, the NRI-binding site and the glnAp2
promoter were inserted into a plasmid vector upstream of a cloning site.
When the lacZ gene was placed under the control
of the glnAp2 promoter, there was no significant βgalactosidase synthesis until late in the exponential
phase of growth, just as expected. A similar result
was obtained when the lacZ gene was replaced
with a construct encoding two different metabolic
enzymes. Finally, as a real test of the system, an engineered construct encoding a pathway for the
synthesis of the carotenoid lycopene was placed
under the control of the glnAp2 promoter. This was
a particularly interesting test because one of the precursors of lycopene, pyruvate, is also an immediate
precursor of acetyl phosphate. Once again, product
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309
Glucose
Glyceraldehyde
3-phosphate
Lycopene
Pyruvate
NRI
Acetyl
phosphate
NRI-P
Acetate
Expression
Fig. 14.20 Construction of a metabolic
control circuit. See text for details.
NRI-P
binding
region
(lycopene) formation was controlled by the metabolic state of the cell. Although metabolic control
engineering is still at an early stage, it represents a
significant degree of sophistication of control compared with the use of simple controllable promoters.
Metabolic engineering in plant cells
Plants synthesize an incredibly diverse array of
useful chemicals. Most are products of secondary
metabolism – that is, biochemical pathways that are
not involved in the synthesis of essential cellular
components but which synthesize more complex
molecules that provide additional functions. Examples
of these functions are attraction of pollinators and
resistance to pests and pathogens. In many cases,
these secondary metabolites have specific and potent
pharmacological properties in humans: well-known
examples include caffeine, nicotine, morphine and
cocaine.
Plants have long been exploited as a source of
pharmaceutical compounds, and a number of species
are cultivated specifically for the purpose of extracting drugs and other valuable molecules. We discussed above how gene transfer to bacteria and
yeast can be used to produce novel chemicals, so in
theory it would be possible to transfer the necessary
components from these useful plants into microbes
for large-scale production. However, the secondary
glnAp2
promoter
lacZ or lycopene biosynthesis
genes
metabolic pathways of plants are so extensive and
complex that, in most cases, such a strategy would
prove impossible. Fortunately, advances in plant
transformation have made it possible to carry out
metabolic engineering in plants themselves, and
large-scale plant cell cultures can be used in the
same manner as microbial cultures for the production of important phytochemicals (reviewed by
Verpoorte 1998, Verpoorte et al. 2000).
The secondary metabolic pathways of most plants
produce the same basic molecular skeletons, but
these are ‘decorated’ with functional groups in a
highly specific way, so that particular compounds
may be found in only one or a few plant species.
Furthermore, such molecules are often produced in
extremely low amounts, so extraction and purification
can be expensive. For example, the Madagascar
periwinkle Catharanthus roseus is the source of two
potent anti-cancer drugs called vinblastine and
vincristine. These terpene indole alkaloids are too
complex to synthesize in the laboratory and there
are no alternative natural sources. In C. roseus, these
molecules are produced in such low amounts that
over 1 ha of plants must be harvested to produce a
single gram of each drug, with a commercial value of
over $1 million.
It would be much more convenient to produce
such drugs in fermenters containing cultured plant
cells, and this has been achieved for a number of
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compounds, two of which (paclitaxel and shikonin)
have reached commercial production (see Verpoorte
et al. 2000). However, cell-suspension cultures often
do not produce the downstream products made by
the parent plant. This applies to vinblastine and vincristine from C. roseus, and also to other important
drugs, such as morphine, codeine and hyoscyamine.
Part of the reason for this is the complexity of secondary metabolism. The entire pathway is not completed in a single cell, but is often segregated into
different cell types, with consequent shuttling of
intermediates between cells. Within the cell, different stages of the pathway are also compartmentalized, so that intermediates must be transported
between organelles. As in bacteria, knowledge of the
target biosynthetic pathway is therefore essential
for metabolic engineering, but in plants only a few
secondary pathways are understood in sufficient
detail. Examples include the phenylpropanoid and
flavonoid pathways, which yield anthocyanins
(plant pigments) and phytoalexins (antimicrobial
compounds), and the terpene indole alkaloid biosynthetic pathway, which generates important alkaloids, such as vinblastine and vincristine. As more
plant genomes are sequenced, we are likely to learn
much more about such pathways.
All terpene indole alkaloids derive from a single
universal precursor called strictosidine. This is
formed by the convergence of two pathways, the iridoid pathway (culminating in secologanin) and the
terpenoid pathway (culminating in tryptamine).
Strictosidine is formed by the condensation of secologanin and tryptamine, catalysed by the enzyme
strictosidine synthase, and is then further modified
in later steps to produce the valuable downstream
alkaloids (Fig. 14.21). The conversion of tryptophan
COOH
L-Tryptophan
NH2
N
H
Tryptophan
decarboxylase
(TDC)
H
O
C
H
OGIu
5
Tryptamine
H
N
H
2
NH2
O
CH3OOC
Secologanin
Strictosidine synthase
(SSS)
N
H H
3
NH
H
OGIu
Strictosidine
H
CH3OOC
O
N
H
N
N
H H
H
CH2
H
CH3OOC
Ajmalicine
O
CH3O
N
H
CH3
Vindoline
OH
OAc
COOCH2
Fig. 14.21 Abbreviated pathway of
terpene indole alkaloid biosynthesis,
showing the conversion of tryptophan
into tryptamine by the enzyme
tryptophan decarboxylase, the
condensation of tryptamine and
secologanin into strictosidine, and the
later diversification of strictosidine into
valuable akaloids, such as ajmalicine
and vindoline (a precursor of both
vinblastine and vincristine).
POGC14 9/11/2001 11:11 AM Page 311
Applications of recombinant DNA technology
to tryptamine is a rate-limiting step in the terpenoid
pathway, and this has been addressed by overexpressing the enzyme tryptophan decarboxylase in
C. roseus cell-suspension cultures. However, while
transformed cultures produced much higher levels
of tryptamine, no downstream alkaloids were synthesized (Goddijn et al. 1995, Canel et al. 1998). The
simultaneous overexpression of the next enzyme in
the pathway, strictosidine synthase, did increase the
levels of the alkaloid ajmalicine, a useful sedative, in
some cultures, but did not result in the synthesis of
vinblastine or vincristine (Canel et al. 1998).
Thus it seems that single-step engineering may
remove known bottlenecks only to reveal the position of the next. The limited success of single-gene
approaches has resulted in the development of
alternative strategies for the coordinated regulation
of entire pathways using transcription factors. Using
the yeast one-hybrid system (Vidal et al. 1996a), a
transcription factor called ORCA2 has been identified that binds to response elements in the genes
for tryptophan decarboxylase, strictosidine synthase
and several other genes encoding enzymes in the
same pathway. A related protein, ORCA3, has been
identified using insertional vectors that activate
genes adjacent to their integration site. By bringing
the expression of such transcription factors under
the control of the experimenter, entire metabolic
pathways could be controlled externally (see review
by Memelink et al. 2000).
Apart from the modification of endogenous
metabolic pathways to produce more (or less) of a
specific endogenous compound, plants can also be
engineered to produce heterologous or entirely novel
molecules. An example is the production of the
alkaloid scopolamine, an anticholinergic drug, in
Atropa belladonna (Hashimoto et al. 1993, Hashimoto
& Yamada 1994). Scopolamine is produced in Hyoscyamus niger but not in A. belladonna, which accumulates the immediate precursor hyoscyamine. H.
niger converts hyoscyamine into scopolamine using
the enzyme hyoscyamine-6-hydroxylase (H6H),
which is absent in A. belladonna. Hashimoto and
colleagues isolated a complementary DNA (cDNA)
encoding H6H from H. niger and expressed it in A.
belladonna. The transgenic A. belladonna plants produced scopolamine because they were able to extend
the metabolic pathway beyond its endogenous
311
end-point. Another example of the production of
novel chemicals in plants is the diversion of carbon
backbones from fatty acid synthesis to the formation
of polyhydroxyalkanoates, which form biodegradable thermoplastics (Steinbuchel & Fuchtenbusch
1998). In this case, the foreign genes derive not from
other plants, but from bacteria.
Theme 6: Plant breeding in the
twenty-first century
Introduction to theme 6
We have discussed several uses for transgenic plants
earlier in this chapter, i.e. as bioreactors producing
recombinant proteins (p. 285) and novel metabolites (see previous section). Transgenic plants can
potentially express any foreign gene, whether that
gene is derived from bacteria, yeast, other plants or
even animals. The scope for exploitation and improvement is virtually limitless, and gene-manipulation
techniques have therefore given the biotechnology
industry a new lease of life. In the following sections,
we consider the development of plant biotechnology
in two key areas: improvement of agronomic traits
and modification of production traits.
Improving agronomic traits
The initial focus of plant biotechnology was on
improving agronomic traits, i.e. the protection of
crops against pests, pathogens and weeds, and thus
increasing yields. Major crop losses are caused every
year by these so-called ‘biotic’ constraints, as well
as physical (or ‘abiotic’) factors, such as flooding,
drought, soil quality, etc. The aims of the biotechnology industry went hand in hand with those of
conventional breeders, but offered the possibility of
importing useful genes from distant species that could
not be used for breeding. It has been found that, in
many cases, single genes transferred from another
organism can provide high levels of protection.
Herbicide resistance
Herbicides generally affect processes that are unique
to plants, e.g. photosynthesis or amino acid biosynthesis (see Table 14.8). Both crops and weeds share
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CHAPTER 14
Table 14.8 Mode of action of herbicides and method of engineering herbicide-resistant plants.
Herbicide
Pathway inhibited
Target enzyme
Basis of engineered resistance to herbicide
Glyphosate
Aromatic amino acid
biosynthesis
5-Enol-pyruvyl shikimate3-phosphate (EPSP)
synthase
Overexpression of plant EPSP gene or introduction of
bacterial glyphosate-resistant aroA gene
Sulphonylurea
Branched-chain amino
acid biosynthesis
Acetolactate synthase
(ALS)
Introduction of resistant ALS gene
Imidazolinones
Branched-chain amino
acid biosynthesis
ALS
Introduction of mutant ALS gene
Phosphinothricin
Glutamine biosynthesis
Glutamine synthetase
Overexpression of glutamine synthetase or introduction
of the bar gene, which detoxifies the herbicide
Atrazine
Photosystem II
QB
Introduction of mutant gene for QB protein or
introduction of gene for glutathione-S-transferase,
which can detoxify atrazines
Bromoxynil
Photosynthesis
these processes, and developing herbicides that are
selective for weeds is very difficult. An alternative
approach is to modify crop plants so that they
become resistant to broad-spectrum herbicides, i.e.
incorporating selectivity into the plant itself rather
than relying on the selectivity of the chemical. Two
approaches to engineering herbicide resistance have
been adopted. In the first, the target molecule in the
cell either is rendered insensitive or is overproduced.
In the second, a pathway that degrades or detoxifies
the herbicide is introduced into the plant. An example of each strategy is considered below.
Glyphosate is a non-selective herbicide that
inhibits 5-enol-pyruvylshikimate-3-phosphate (EPSP)
synthase, a key enzyme in the biosynthesis of
aromatic amino acids in plants and bacteria. A
glyphosate-tolerant Petunia hybrida cell line obtained
after selection for glyphosate resistance was found to
overproduce the EPSP synthase as a result of gene
amplification. A gene encoding the enzyme was subsequently isolated and introduced into petunia plants
under the control of a cauliflower mosaic virus
(CaMV) 35S promoter. Transgenic plants expressed
increased levels of EPSP synthase in their chloroplasts and were significantly more tolerant to
glyphosate (Shah et al. 1986). An alternative approach
Introduction of nitrilase gene, which detoxifies
bromoxynil
to glyphosate resistance has been to introduce a
gene encoding a mutant EPSP synthase. This
mutant enzyme retains its specific activity but has
decreased affinity for the herbicide. Transgenic
tomato plants expressing this gene under the control
of an opine promoter were also glyphosate-tolerant
(Comai et al. 1985). Following on from this early
research, several companies have introduced glyphosate tolerance into a range of crop species, with
soybean and cotton the first to reach commercialization (Nida et al. 1996, Padgette et al. 1996). Currently,
nearly three-quarters of all transgenic plants in the
world are resistant to glyphosate (James 2000).
Phosphinothricin (PPT) is an irreversible inhibitor
of glutamine synthetase in plants and bacteria.
Bialaphos, produced by Streptomyces hygroscopicus,
consists of PPT and two alanine residues. When
these residues are removed by peptidases the herbicidal component PPT is released. To prevent selfinhibition of growth, bialaphos-producing strains
of S. hygroscopicus also produce the enzyme phosphinothricin acetyltransferas